Objectives: Kidney transplant in mouse model is quite useful and valuable for studying transplant immunobiology. However, its technical complexity and high mortality have hindered widespread use. We sought to review and analyze the various techniques of kidney transplant in mice to prevent pitfalls and reduce complications.
Materials and Methods: We searched PubMed using the key words “kidney transplant” or “renal transplant,” “mouse” or “mice,” and “technique” or “model” for articles published between January 1973 and June 2010. We found a series of factors that were collected and analyzed, which may influence establishing a mouse kidney transplant model.
Results: A mouse kidney transplant model is practical for research. Mouse body weight, sex, animal strain, type of anesthesia, and type of flushing solution are relevant but do not determine a successful transplant. Warm and cold ischemia time should be less than 35 minutes and 2.5 hours.
Conclusions: End-to-side vascular anastomosis and ureter-to-bladder or bladder-to-bladder for ureter reconstruction is preferred for establishing a kidney transplant model in mice.
Key words : Kidney transplant, Mouse model, Strategies, Influencing factors, Techniques
The mouse model has been used for research since there are more than 1300 mutant mice with which to study the biology of diseases, particularly the complexity of the immune system.1 Indeed, the mouse has many inherent advantages: low cost, no specialized animal care, short gestation, short life spans, and no culture sensitivities. In the field of solid organ transplant, use of various mice of genetically well-defined inbred strains, congenic and transgenic strains, has become an instructive and effective strategy to clarify pathological conditions or molecular pathways in genetics, immunology, and nonimmunology including ischemia/reperfusion.2-8 In particular, investigating the mouse H-2 system will improve our understanding of human leukocyte antigen complex based on their high-level similarity.9-10 Furthermore, developing multiple biological technologies (such as monoclonal antibodies and molecular biology probes) provide wide availability for the mouse rather than other laboratory animals.11 Therefore, the mouse transplant model can, in some measure, mimic its human counterpart, providing mechanistic insight into clinical settings.
The mouse model of a kidney transplant was first reported in 1973 by Skoskiewicz and associates.12 Different from other transplant models, kidney-recipient mice engrafted with major histocompatibility complex-mismatched donors can survive for 54 days without treatment,4 while additional administration of anti-CD154 monoclonal antibodies can lead to long-term graft survival (> 100 days)4 across major histocompatibility barriers. However, the demanding techniques and mortality rates have impeded widespread use of this model. To better understand the difficulties of mice kidney transplant, we reviewed the techniques for a mouse kidney transplant and evaluated the advantages and disadvantages of different methods according to 3 determinants: Requirement for microsurgical skill, speed of reconstruction, and closeness to physiological setting.
Materials and Methods
Data were collected through a MEDLINE database using the key words “kidney transplantation” or “renal transplantation,” “mouse” or “mice,” and “technique” or “model.” The search identified 7 articles that discussed surgical techniques, while other articles related to mouse kidney transplant were carefully reviewed. Mouse kidney transplant is demanding and warrants meticulous technique. All modifications were made to sew the vessels rapidly and easily, to reduce ischemia time, and to improve recipient survival.13 Therefore, a series of important factors probably influencing establishment of a mouse kidney transplant model were collected and analyzed.
Selection of animals
Inbred mouse strains are usually used for kidney transplant, irrespective of the different aims of study. Congenic mouse strains are genetically identical, except for major histocompatibility complex allele (class I and/or class II). The major histocompatibility complex-disparity between donor and recipient induces an allogeneic response, leading to various graft survivals. Based on current research investigations, C57BL/6, BALB/c, and their derived strains (including gene knockout strains) are a major source of mice used for transplant studies,7, 11, 14-19 in addition to CBA/J,15 C3H,20 and DBA/2 mice.6, 16, 19 Transplant using different combinations of these mice can elicit disparate immune responses, which meet a need for short-, medium-, long-term, or third-party survival studies.
Either clinical21 or rat22 experimental investigations show that age is a nonimmunological determinant influencing graft function. To some extent, rat body weight is directly proportional to its age,23 while mouse age is not linearly correlated with weight. In previous experimental transplant models, mice 6 to 12 weeks old were used.5, 15, 24 Nevertheless, all articles11, 13, 14, 17, 20, 25 underscored a weight of 20 to 30 grams in mice destined for surgery; except Han and associates, who used mice that weighed less than 20 grams.15 Interestingly, heavier mice may have extensive fatty tissue without having larger caliber vessels, which may not be beneficial for the vessels’ anastomosis.20
Not only clinical26 but also experimental data27 show that sex plays an important role in transplant. Male mice were used mainly for mouse kidney transplant11, 13, 20, 25; however, females were used in 2 studies.15, 16 The predominant selection of male mice was made because ureter reconstruction by means of bladder-bladder anastomosis is simpler in males14 and hormonal changes do not exist. In addition, the penile vein can be used for intravenous injection, especially for large-size cell transfer. Conversely, the female uterus occupies the space for reconstruction of urinary drainage. Experimental investigations show no detectable differences in the response of a female recipient receiving a transplant from a female donor.14
General anesthetics composed of inhalation and injection (intramuscular [IM] or intraperitoneal [IP]) are normally given to small animal surgeries. Special equipment for inhalation and injection was adopted as an inexpensive approach to anesthetize mice5, 11, 14, 15, 20, 25 (eg, combined use of ketamine and xylazine [IP], chloral hydrate, and methoxyflurane [IP], 4% halothane). Although these do not affect the transplant outcome, caution should be exercised to avoid complications.
Most researchers use the left donor kidney for transplant, as the renal vessels of the right kidney are not long enough for reconstruction.12, 14 Nonetheless, Kalina and associates successfully conducted mice kidney transplants using both the left and right donor kidney with equal success.13
Donor preservation and heparinization
There are 2 kinds of heparinization for the donor: Kidney graft heparinization and somatic heparinization. Although somatic heparinization (100 U/mL) is not indispensable, its use to avoid thrombosis and ensure graft perfusion is well documented. Most studies use the donor kidney, perfused slowly with cold heparinized solution (100 U/mL) in situ or via the infrarenal aorta. Albeit no literature has described the maximal dosage of heparin for perfusion, our experience shows that 100 U/mL is optimal to avoid delayed hemorrhage, and the direct use of the original concentration (10 000 U/mL) causes anastomotic bleeding and even death of the recipient. Indeed, an extreme loss of blood volume, despite survival of the transplant recipient, can engender hematocytopenia and reactive lymphoproliferation, affecting subsequent immune analysis and data reliability. Direct perfusion of the kidney graft may minimize blood transfer from the donor.13 Perfusate, including blood, was drained via the cut end of the inferior vena cava.11, 15, 25 Additionally, Nesbitt and associates have applied heparin, not only to the donor, but to the recipient, at the concentration of 1.5%.5 Care must be taken to avoid overheparinization, inducing a delayed hemorrhage.
Different perfusion solutions have been suggested, such as phosphate buffered saline (4°C),17 normal saline,15, 25 and lactated Ringer’s solution.11, 14, 19, 20 The en bloc kidney was freed, removed, and placed in a cold solution. The preservation solution may be the same with a perfusion solution,17 normal saline,15 lactated Ringer’s,20 or a different solution (eg, University of Wisconsin or histidine-tryptophan-ketoglutarate solution). The latter is preferable when the transplant will be performed more than 30 minutes after procurement.25 The addition of heparin (5 U/mL) in the preservation solution prevents clot formation.5 Owing to the protective role of the hypertonic solution in the graft, mannitol, and Chloromycetin sodium also have been considered for use in the preservation solution.5
Cold ischemia starts from kidney perfusion, through procurement, and goes until the end of cold storage at 4°C. Since the initial attempts of perfusing an isolated organ by Loebel in 1849,28 the deleterious effects of cold ischemia time have been widely studied in human and rat, but not as much in the mouse. Farrar and associates reported that prolonged cold ischemia (2 hours) could result in a significant increased expression of intrarenal C3 protein, an acute-phase protein induced by cold-ischemia injury in several organ models.29 Reduced renal graft function has been observed at early time points (2.5~3 h).30 Furthermore, histologic analysis shows renal tubular injury becomes severe, as cold ischemia reaches 4 hours in the mouse model.29
Warm ischemia refers to the period from positioning the graft in the recipient until reconstruction of graft blood flow. In the course of the vessels’ reconstruction, microscopy lamplight and body temperature of the recipient may increase the chilled graft’s temperature, aggravating the graft injury. Therefore, cold normal saline-soaked gauge is suggested for wrapping the kidney graft. In the rat kidney transplant model, previous investigations required warm ischemia of less than 45 minutes.31 In general, to achieve better graft survival, particularly with long-term graft outcome, fewer than 35 minutes of warm ischemia was suggested in a mouse model based on previous studies.11, 13, 14, 17, 20, 25
Preparation for vascular anastomosis
The diameter (0.4 mm) of mouse vascular lumen for anastomosis is 20 times smaller than that of a rat,11 which requires meticulous microsurgical manipulations. The vascular intima is easily impaired during the aorta and inferior vena cava of recipient, which may result in later thrombus formation. The aorta and inferior vena cava between the renal vessel and iliac bifurcations are freed and exposed (approximately 1 cm) for anastomosis. To block blood flow and distend the vessels, the proximal and distal ends of these vessels may be placed with 4-0 silk ties,13, 14 or 7-0 silk ties,11, 25 or (4 mm) microvascular clamps.11, 20
A few lumbar branches should be cauterized11 or ligated14, 20, 25 before aortotomy and venotomy. If continuous blood filling is observed in the IV after cross-clamping, ligation of the lumbar branches or microvascular clamps should be checked. Our experience has proved that cauterization is feasible, saving time when compared with ligation.
An elliptical aortotomy the diameter of approximately one-fifth of the aorta is made using an insulin syringe with a 0.36-mm Microfine needle13 or by cutting with an iris scissors under the guidance of an 11-0 suture in the aorta.20 It is crucial to penetrate the full thickness of aorta, which reduces risk of thrombosis or pseudoaneurysm formation. A larger opening should be avoided; otherwise, it could lead to anastomotic constriction and subsequent thrombosis.20
A venotomy is made longitudinally in the inferior vena cava with a 30-gauge needle and pruned with an iris scissors. The size of the opening matches the diameter of the donor’s inferior vena cava. Flushing thoroughly with heparinized saline is necessary to clean intraluminal blood clots and prevent thrombosis after surgery.20
With a view of the renal vessels in mouse, the aorta and inferior vena cava must be used for vessels’ reconstruction, strictly recognized as heterotopic transplant instead of orthotopic.18, 20 One exception is Lutz and associates who reported a de facto orthotopic kidney transplant in mice.17 Unlike a rat kidney transplant, it is crucial to first prepare the recipient including exposing the inferior vena cava and aorta for reconstruction before donor organ procurement, which will minimize cold ischemia of kidney graft, reduce the risk of postoperative paraplegia of recipient, and increase the success rate of surgery.11, 13 Using the donor suprarenal aorta and inferior vena cava for a vascular anastomosis avoids the difficult dissection that renal vessels require and reduces the negative influence of the blood supply of the donor graft during explantation and ensures the quality of the procured kidney graft and speed of reconstruction.13
Besides an end-to-end anastomosis, end-to-side anastomosis of the donor’s and the recipient’s aortas or inferior vena cava is frequently done with 10-0 or 11-0 nylon. A continuous suture is consistently adopted by all authors for vascular anastomosis (Table 1). Care should be taken to orient the donor aorta and avoid twisting the inferior vena cava during anastomotic surgery. Arterial reconstruction is done first. As few stitches as possible are used for arterial anastomosis by atraumatically manipulations, minimizing the risk of thrombosis.20 In view of thin and fragile venous wall, saline irrigation is helpful to visualize the openings during anastomotic suturing. At least 4 stitches on each side should be performed for venous reconstruction.11, 20
A bit of Gelfoam or microfibrillar collagen will minimize bleeding by placing it around the anastomotic sites before ties or microvascular clamps are released. Afterwards, suitable pressure with a dry swab is applied for a while.13, 20
Urinary tract reconstruction
Ureter anastomosis is also a challenging part of kidney transplant in a mouse model. Different urinary complications frequently are seen depending on specific reconstruction approaches such as necrosis, urine leakage, ureter constriction or obstruction, or hematuria, despite a lack of calculi formation. Unlike a rat model, no available stent or cannula is reported in the mouse owing to its tiny and delicate ureter. Correct orientation of the ureter is required to avoid kinking.
This approach includes bladder patch-to-patch13, 20 and bladder patch-to-cystotomy.25 With respect to the former, as the dome of the recipient bladder is dissected, bleeding may be controlled by cautery20 or by using a 4-0 silk tie around the midportion of bladder.13 The donor bladder patch without sufficient blood supply should be small to avoid necrosis. And the size of the recipient’s bladder dome should be tailored to fit. The anastomosis is usually done with interrupted or continuous 9-0 or 10-0 nylon sutures (Table 1).14, 20 The method of bladder patch-to-cystotomy has been reported by Martins, which was developed from the Lich-Gregoir technique for extravesical ureteroneocystostomy.25 At the time of implantation, the recipient’s bladder is usually filled with urine, facilitating performance of the cystostomy. Despite its advantage of antireflux, the technique is time consuming and more complicated.
For ureter reconstruction, the technique of a ureter-bladder anastomosis is preferred. The bladder is perforated with a 20- or 21-gauge needle.11, 15 The distal end of the ureter is pulled into the bladder. Although 2 stitches are needed to fix the ureter to the wall of the bladder, the implanted ureter still might escape from the bladder. Therefore, it is necessary to position the kidney graft close to recipient’s bladder to reduce the tension of implanting the ureter into bladder.15 Although it is important to keep the periureteric tissue and avoid harming the ureter’s blood supply, the end of the periureteral nourishing vessels should be cauterized or ligated to prevent hematuria. In our experience, improper management of these vessels frequently leads to hemorrhage of the urethra.
Despite the technical simplicity of ureter-to-bladder anastomosis, we found that different the degree of hydronephrosis (mostly mild) was observed in long-term surviving grafts (even syngeneic grafts) and accounted for compression of the ureter by the bladder or by vesicoureteral reflux. Severe hydronephrosis can end graft survival. By contrast, given that bladder-to-bladder anastomosis is adopted, urine reflux from the bladder back to the kidney can be avoided. Conversely, by using a ureter-to-bladder anastomosis, if the size of the perforated hole of the bladder is much bigger than the inserted ureter, urine leakage may occur. Based on our experience, mild hydronephrosis is acceptable, even for long-term survival studies, and histology staining usually shows normal kidney signs in the early stage posttransplant (Figure 1). Periureteral connective tissues can be used and sutured to the outside of the bladder hole to prevent urine leakage.
End-to-end ureter anastomosis
Unlike a rat kidney transplant, the diameter of the mouse ureter is tiny (< 0.4 mm); therefore, placement of polyethylene stent into a mouse ureter or for cannulation is unfeasible for ureter reconstruction. Only Lutz and associates reported this technique of ureter connection by end-to-end anastomosis using 10-0 Prolene sutures. But no more details were shown.17
Postoperatively, urine is normally produced in approximately 3 minutes. No urine production is probably caused by ureter blockade, thrombosis of renal vessels, or acute tubular necrosis.
To evaluate the function of graft precisely, native kidneys of recipient are normally removed. One of them is consistently replaced by graft at the time of implantation. Contralateral native kidneys may be resected concurrently13, 14, 20 or at day three,4, 18 day four,5-7 day five,19 day seven,20, 32, 33 or day ten17 after transplant. By comparison, 2-stage procedure that native kidneys are excised at different times may significantly increase the success rate of transplant surgery,20, 32 as evidenced by an experiment that transplanted isograft with bilateral nephrectomy tended to uremia with 36 hours.14
Running stitch with absorbable 4-0 or 5-0 silk was applied to close the abdominal wall. Skin and muscle may be sutured in a single layer13 or separately.5, 25
Fluid replacement and postoperative care
Owing to the larger ratio of surface area/body weight compared with other laboratory animals, body temperature of mice is prone to decrease after surgery. For the first 24 hours postoperatively, animals wrapped in gauze are normally kept on a heating pad (37°C)11 or under a 100-watt heating lamp13, 20, 25, 34 to maintain body temperature. In addition, the use of circulating water blankets is recommended to avoid severe burns or hyperthermia if the mouse is anesthetized or unable to escape from the heat lamps or heating pads. Mice usually recovered from anesthesia within 1 hour after the operation and are given regular food and water ad libitum.20
To investigate immunobiological mechanism, various knock-out mice are normally adopted, whose immune system is defective and susceptible to viral and bacterial infection. Although sterile surgery is required for transplant, antibiotics are normally administered to recipients to avoid infectious complications, particularly for the first 24 hours posttransplant. Cephtriaxone11 and penicillin (500 U/10 g)20, 33 were reportedly used. In addition, co-trimoxazole may be added in the drinking water.33 However, Martins did not suggest use of any antibiotics after transplant.25
Kidney transplant surgery is a relatively large surgery for mouse. Intraoperative blood loss is inevitable. Long-time operations together with heat from a microscopic lamplight may cause over-exertion and apparent body fluid loss. Therefore, during or after surgery, the recipient is usually supplied with warm physiological saline subcutaneously and/or via vein injection. The amount varies widely,11, 20 depending on blood loss, total operation time, and intensity of lamplight; 0.5 mL normal saline solution may be injected subcutaneously.25
Daily observation of recipient health status is important to assess kidney graft function. After 2 native kidneys are removed, the kidney graft may be directly evaluated by recipient survival. Kidney function is monitored by determining serial blood urea nitrogen levels and serum creatinine levels.11, 13, 14, 19 Blood may be required from the retro-orbital plexus14 or the tail vein. Normal range of serum creatinine level is 16 to 28 µmol/L.15, 19 Elevation of serum creatinine (~100 µmol/L) and signs of illness may represent kidney graft rejection.19 Normal range of blood urea nitrogen is 0.15 to 0.30 mg/mL with an average of 0.224 mg/mL.14 To accurately assess kidney graft function, phosphate clearance by infusion of [14C]inulin via the left jugular vein is suggested.5 Considering technical restriction, serial biopsies of the transplanted graft cannot be fulfilled in practice. The harvested graft can be analyzed by histopathology.14
Success of the operation
The success rate reported before differs greatly between 50% and 90%.12, 13, 15, 20, 32 Although sometimes the microsurgeon has much experience with other transplant models, with mice heart and pancreatic transplants only 50% survived.12
Admission standard of transplanted recipients
Bilaterally nephrectomized animals that do not survive more than 3 days should be considered a technical failure, regardless of cause.14
Although the mouse model of kidney transplant has been established for over 30 years, the complicated and challenging techniques hamper its widespread use. To improve mastery of these techniques, some contributing factors should be determined.
Perfusion and preservation solution
To procure a high-quality donor graft, University of Wisconsin solution is recommended for use, particularly as the graft is harvested with prolonged ischemic time.
Avoidance of complications and frequent technical failures
For the beginner, unreasonable anesthesia induction may cause animal death. Mishandling of renal vessels may impair the graft during organ procurement. Apart from these, vascular thrombosis, uncontrolled bleeding, anastomotic constriction, urine leakage, intrabladder hemorrhage, obstruction of the urinary system, and paraplegia are frequently complications that may develop within 24 hours posttransplant.32 Death owing to intraoperative infections or hypovolemia may occur even when 1 native kidney is kept in the mice.
Avoidance of technical errors and minimize blood loss
Successfully establishing a kidney transplant in the mouse requires meticulous and delicate dissection.13 Excessive manipulations lead to vasospasm and vessel injury, leading to thrombosis during recipient surgery. Minimizing blood loss less than 0.1 mL is critical to improve the success rate of long-term graft survival.32
Reduction of cold/warm ischemic injury
Warm ischemic time under 35 minutes is advantageous to graft outcome. However, it does not mean the shorter time of warm ischemia will consequentially lead to better graft survival. Although the graft is protected by cold moisturized gauze against warm ischemic injury, the graft may benefit from a rinse with cold saline several times during the anastomosis.11, 20 Prolonged cold ischemic time may lead to a poor graft outcome, reducing the long-term survival of recipient.
Technique of vascular reconstruction
The end-to-side procedure is widely used for establishing a mouse model of kidney transplant. Continuous sutures are preferred for vascular anastomosis with high success, to our knowledge, despite the fact that interrupted suture is possible to successfully complete artery reconstruction, which will prolong warm ischemic time.
Technique of urinary tract reconstruction
By using a bladder-to-bladder anastomosis, care should be taken to keep the sufficient blood supply for the donor bladder patch. Otherwise, ischemia, even necrosis, would occur after a bladder-to-bladder anastomosis. With respect to this aspect, it usually takes at least 30 minutes more for the donor and recipient surgeries, as compared with the technique of implanting ureter into bladder.
In conclusion, the techniques provided in this review article offer various options for learners. The most viable should be used for practice. Care for some important factors may avoid operative complications, improve surgical techniques, and shorten the learning curve.
Volume : 9
Issue : 5
Pages : 287 - 294
From the 1Department of Administration and Development, the 2nd Affiliated
Hospital of School of Medicine, Zhejiang University, Hangzhou City, People's
Republic of China; 2Department of Public Health, Charité, Campus Virchow Clinic,
Universitätsmedizin Berlin, Germany; and the 3Departments of Medicine and
Surgery, Harvard Medical School, Transplant Institute, Division of Immunology,
Beth Israel Deaconess Medical Center, Boston, MA 02215, USA
Address reprint requests to: Dr. Weihua Gong, CLS 617, 3 Blackfan Circle, Boston, MA 02215, USA
Phone: +1 617 806 6889
Fax: +1 617 735 2902
Table 1. Various techniques for reconstruction of renal vessels and ureter in mice kidney transplant.
Figure 1. Histologic analysis of syngeneic graft.
Syngeneic graft harvested from wild- type C57BL/6 on day 7 posttransplant was snap frozen into Tissue Tec and staining with Harris Hematoxylin and Eosin, showing normal proximal tubule (asterisks), normal Bowman’s capsule (arrows) and normal glomerulus (× 250 of original magnification).