Here we present a murine model of arteriovenous fistula (AVF) failure in which a clinically relevant anastomotic configuration is incorporated. This model can be used to study the pathophysiology and to test possible therapeutic interventions.
The arteriovenous fistula (AVF) still suffers from a high number of failures caused by insufficient remodeling and intimal hyperplasia from which the exact pathophysiology remains unknown. In order to unravel the pathophysiology a murine model of AVF-failure was developed in which the configuration of the anastomosis resembles the preferred situation in the clinical setting. A model was described in which an AVF is created by connecting the venous end of the branch of the external jugular vein to the side of the common carotid artery using interrupted sutures. At a histological level, we observed progressive stenotic intimal lesions in the venous outflow tract that is also seen in failed human AVFs. Although this procedure can be technically challenging due to the small dimensions of the animal, we were able to achieve a surgical success rate of 97% after sufficient training. The key advantage of a murine model is the availability of transgenic animals. In view of the different proposed mechanisms that are responsible for AVF failure, disabling genes that might play a role in vascular remodeling can help us to unravel the complex pathophysiology of AVF failure.
A functional vascular access conduit is of vital importance for patients with renal failure that depend on chronic hemodialysis to stay alive. The construction of an arteriovenous fistula (AVF) is currently the preferred choice for vascular access. However, AVF related complications constitute a major cause of morbidity for patients on chronic hemodialysis. Despite extensive scientific efforts, none of the novel approaches to reduce AVF access-related complications did result in substantial improvement of AVF durability. Part of this disappointing progress relates to incomplete understanding of the underlying pathophysiology of hemodialysis access failure.
To unravel the pathophysiology of AV access failure, animal models that closely mimic human pathology are of utmost importance. In this respect, not only the animal species but also the anastomotic site, the required anti-coagulatory therapy and the duration of follow up after surgery should be taken into account1. While large animals are the most suitable for intervention studies aimed to develop new therapeutic strategies, murine models have the greatest potential to gain more insight in the molecular mechanisms underlying AV access failure due to the availability of transgenic mice. In addition, a large number of mice can be used for this purpose at lower costs compared to larger animal use.
The first murine model of AVF failure was described in 2004 by Kwei and et al.2 In this model, AVFs were constructed using the carotid artery and the jugular vein in an end-to-end manner using an intravascular catheter. This model could be useful to study early venous adaptation in AVFs although the end-to-end configuration and the presence of an intravascular catheter limit the validity of this model for human AVFs. An improved AVF model was introduced by Castier and et al.3 in which the end of the carotid artery is connected to the side of the jugular vein. However, AVFs in hemodialysis patients are usually constructed by anatomizing the end of a vein to the side of an artery. The exact configuration of the AVF is a crucial characteristic of an AV access model since it determines the hemodynamic profile within the conduit4. The latter is an important contributor to endothelial dysfunction and subsequent development of intimal hyperplasia (IH)5.
A novel murine model was recently developed with an identical anatomical configuration as is utilized in humans6. In this model, AVF are created in C57BL/6 mice by anastomosing the end of a branch of the external jugular vein to the side of the common carotid artery with interrupted sutures. In the present paper, we focus on the microsurgical procedure of this model in order to facilitate the widespread use of this murine model, aimed to unravel the complex pathophysiology of hemodialysis access failure.
All experiments were approved by the committee on animal welfare of the Leiden University Medical Center.
1. Animal Preparation and Anaesthesia
2. Skin Incision
3. Dissecting and Preparing the Vein
4. Removing the Sternocleidomastoid Muscle
5. Dissecting and Preparing the Common Carotid Artery
6. Ligation of the Vein
7. Creation of Anastomosis Part 1
8. Heparin Administration
9. Creation of Anastomosis Part 2
10. Vascular Clamp Removal
11. Skin Closure and Postoperative Care
12. Tissue Harvesting
13. Tissue Embedding and Sectioning
After the creation of the anastomosis (Figure 1), the patency should be assessed by shortly occluding the venous outflow tract with a vascular forceps. When the anastomosis is patent, the vascular tract proximal to the occlusion should clearly expand in a pulsatile manner. In addition, the patency is confirmed by using near infrared fluoroscopy (NIRF) that effectively functions as an angiography (Figure 2). A failure in the surgical procedure can lead to an occlusion of the anastomosis as is depicted in (Figure 2). This failure can be caused by a too narrow anastomotic area, torsion of the vessels, inadequate heparin dose or an accidental suture placement that connects the front side of the anastomosis to the back side.
At a histological level, the process of vascular remodeling in AVF can be investigated elegantly using this model. Vascular remodeling in AVF occurs as a result of the increase in blood flow and pressure. In mice, this response is characterized by an increase in circumference (e.g., outward remodeling) leading to an increase in luminal area in the first 2 weeks after surgery. After these 2 weeks, the luminal area progressively decreases due to a stop in the outward remodeling and ongoing thickening of the tunica intima. The formation of these progressive stenotic lesions results in occlusion of 50% of AVF at 4 weeks after surgery. Therefore, the optimal time point to harvest the AVFs would be at 2 weeks after surgery, since in this phase, proper analysis of the vascular response in patent AVF is still feasible. Immunostaining of the venous outflow tract at 2 weeks after surgery shows that the cellular compartment of the intima mainly consists of alpha-smooth muscle actin (α-SMA) positive cells (Figure 3), as observed in failed human AVFs as well7.
In view of the complexity of the microsurgical procedure, it's realistic to reckon with technical failure of the procedure in a proportion of the mice. In our hands, the success rate of the procedure was 67% in the beginning. However, with further training this rate was increased up to 97%. The main cause of failure was hemorrhage (60%), followed by acute thrombosis (27%) and anesthesia- related death (13%). After sufficient training, the surgical procedure can be performed in approximately 1 hr.
Figure 1. Detailed scheme of surgical procedure. (A–T) Key steps for successful creation of an AVF. Please click here to view a larger version of this figure.
Figure 2. Macroscopic pictures of patent and occluded AVFs. (A) A patent AVF with a vascular clamp on the distal common carotid artery. (B) An occluded AVF. (White arrow) indicates the direction of the blood flow. (C) A patent AVF demonstrated using Near Infrared Fluoroscopy. (red arrow) indicates artery. (blue arrow) indicates venous outflow tract. Please click here to view a larger version of this figure.
Figure 3. Histological stainings of the venous outflow tract of the AVF at day 14 versus unoperated control vessels. (A–B) Morphologic overview using hematoxylin, phloxin and saffron showing a clear increase in vessel circumference and intimal hyperplasia development 14 days after AVF creation. (C–D) Immunohistochemical staining demonstrating that the majority of the cells present the intimal hyperplasia are alpha smooth muscle actin positive. (L) Lumen; (IH) Intimal hyperplasia; scale bar: 200 µm. Please click here to view a larger version of this figure.
The AVF is considered to be the Achilles' heel in hemodialysis treatment. Unfortunately, the AVF still suffers from a high number of failure8-10. Despite extensive research on the underlying mechanisms, the exact pathophysiology remains unknown. Numerous murine models for AVF failure have already been described in literature2,3,11,12. However, none of these models incorporate a venous end to arterial side anastomosis configuration that is most used in the clinical situation. This is very relevant since the resultant hemodynamic profile plays an important role in vascular remodeling. Moreover, some of the models used a synthetic cuff for the connection between the artery and vein2,11, which is not used in the clinical setting.
To improve the clinical relevance, we therefore developed a murine model in which a unilateral venous end to arterial side anastomosis was created between a branch of the jugular vein and common carotid artery with interrupted sutures6. In this model crucial histomorphological changes were observed including outward remodeling and progressive intimal hyperplasia, ultimately leading to AVF failure.
A crucial aspect of the surgical procedure is the anaesthesia protocol. It is recommended to use isoflurane inhalation anaesthesia, as this is a safe and easy method for obtaining prolonged periods of anaesthesia. The latter is especially important for the training phase in which the whole procedure can take up to 3 hr.
In order to create the necessary room for the traversing outflow vein of the arteriovenous fistula, the ipsilateral sternocleidomastoid muscle needs to be excised.
With respect to the vessel handling during surgery, dissection of vessels should be performed in a blunt manner using forceps. The use of specialised vascular straight forceps are recommended for direct handling and manipulation of the vessels as they provide more precision and produce less mechanical damage due to the rounded tip. The usage of suture threads as vessel loops combined with a hemostat can aid in safe tissue handling and moreover, in presenting the tissue in an optimal manner when used as a third hand which is crucial so this procedure can be performed without any direct assistance.
Undoubtedly, the most difficult step in the procedure are the suture placements between the artery and vein. Care must be taken not to suture the "back"-side of the anastomosis with the "front"-side, as this will lead to narrowing of the anastomosis that can lead to early failure of the AVF. To prevent this from happening, position the tip of the vascular forceps between the two walls of the vessel in order to separate them. The technical difficulty can be considered to be a limitation of this model. However, we do not think that this procedure is more challenging than the model that is presently most widely used3,6.
It's difficult to make a general remark on the sample size for future studies, as the calculated sample size does not only depend on the variability between animals but also on the 'strength' of the intervention. We estimate that a group size of approximately 10 mice (excluding mice that drop out of the study because of technical failure of the AVF) should be sufficient for studies that focus on the role of a specific gene in AVF failure. Recently, we were able to obtain significant results with an average group size of 10 animals in a study on the role of elastin in murine AVF remodeling13.
One aspect of our murine model requires further discussion. It is known that the uremic milieu in patients with end stage renal disease contributes to a multitude of vascular diseases including venous intimal hyperplasia even prior to hemodialysis access surgery14-16. This present model does not incorporate the uremic milieu. Therefore, the incorporation of a model of chronic renal failure17 would be a valuable additive step to further improve the validity of this murine model.
The authors have nothing to disclose.
This study was supported by a grant from the Dutch Kidney Foundation (KJPB 08.0003).
Carolien Rothuizen is acknowledged for her contribution to the study. Hoang Pham is acknowledged for his assistance with the pathology work-up.
Dissecting microsocpe | Leica | M80 | |
Forceps | Medicon | 07.61.25 | |
Vascular forceps | S&T | JFL-3D.2 | |
Vascular forceps | S&T | D-5a.2 | |
Forceps | Roboz | SS/45 | |
Micro scissor 5 mm blade | Fine science tools | 15000-08 | |
Micro scissor 2 mm blade | Fine science tools | 15000-03 | |
Scissor | Medicon | 05.12.21 | |
Clip applier 1 | S&T | CAF-4 | |
Vascular clamp 1 | S&T | B-1V | |
Clip applier 2 | BBraun | FE572K | |
Vascular clamp 2 | BBraun | FE740K | |
Hemostatic forceps | BBraun | BH110 | |
10.0 sutures | BBraun | G1117041 | |
6.0 sutures | BBraun | 768464 | |
Cauterizer | Fine science tools | 18010-00 | |
Needle holder | Medicon | 11.82.18 | |
Ocular ointment | Pharmachemie | 41821101 | |
Chlorhexidine tincture 0,5% | Leiden University Medical Center | NA | |
Heparin | Leo Pharma | 012866-08 | |
Buprenorphin | RB Pharmaceuticals | 283732 | |
Isoflurane | Pharmachemie | 45,112,110 | |
Anesthesia mask | Maastricht university | custom made | |
Midazolam | Actavis | AAAC6877 | |
Dexmedetomidine | Orion | 141-267 | |
Fentanyl | Bipharma | 15923002 | |
Continuous anaesthetic induction chamber | Vet-tech solutions | AN010R |