Renal transplantation has become an accepted treatment for end-stage renal failure in cats. However, renal transplantation in cats has unique challenges with respect to the inherent difficulty of the surgical techniques and perioperative management required to maximize the potential for survival of both the graft and patient. Protocols and techniques used for renal transplantation in veterinary species have continued to evolve, particularly in the last decade, as more institutions have gained experience with the procedure.
In human medicine, donated kidneys (grafts) are frequently obtained from brain-dead patients who are located some distance from a regional transplant center1; therefore, the grafts routinely undergo ex vivo CS to maintain their viability during the time required for tissue typing and extensive donor-recipient matching analyses and transport to the facility where the transplantation into the recipient will occur. Hypothermic machine perfusion of ex vivo kidneys is also used in human medicine to help minimize delayed graft function and increase graft survival.2 Ex vivo CS of donated kidneys was not performed when renal transplantation was first developed in cats because the grafts were generally obtained from unrelated but allogeneic living donors and all donor-recipient matching analyses were performed prior to graft harvest.3 Furthermore, donor and recipient operations were frequently performed simultaneously at the same facility and prolonged ex vivo storage of the graft was not necessary.
The approach to transplantation in which the donor and recipient undergo simultaneous surgeries (IT) results in the graft being exposed to warm ischemia for a finite period from clamping of the renal vasculature and removal of the kidney from the donor to completion of vascular anastomoses in the recipient and removal of the vascular clamps. Some degree of renal injury, including the potential for acute or chronic dysfunction, is expected to result from that period of ischemia. Ischemic injury is minimized by limiting the time between clamping the renal vasculature in the donor and removing the clamps following vascular anastomoses in the recipient to preferably < 1 hour, a goal that pressures surgeons to hasten an already demanding technical procedure. Currently, it is unclear to what extent that ischemia contributes to slow or delayed graft function in recipient cats. Results of studies4,5 involving cats that underwent experimental renal transplantation by use of IT without CS indicate that the procedure results in some extent of acute graft dysfunction, which is typically characterized by abnormally increased serum creatinine concentrations after surgery. However, the complexity of the transplantation procedure makes it difficult to determine whether the impaired graft function is a direct result of ischemic injury or associated with technical problems inherent to the procedure. Clinical descriptions of CS methods for feline kidneys are limited, although results of some studies4–6 suggest that flushing ex vivo kidneys with a sucrose phosphate solution followed by CS resulted in peak posttransplantation serum creatine concentrations in recipient cats that were equivalent to or substantially less than those in cats that received a kidney by use of the IT procedure (ie, no CS) even when CS lasted up to 3 hours, which is substantially longer than the period of warm ischemia during the IT procedure.
In human medicine, CS of grafts has been routinely used to suppress warm ischemic injury for nearly 40 years.7 Most mechanisms of warm ischemic injury are mediated by enzymatic processes, and consequently, the rate of activity for those mechanisms is temperature-dependent. Cold temperatures have been used to slow the rate of cumulative ischemic cell damage and extend the period of time that grafts can be feasibly stored ex vivo with minimal injury. Cold storage is an effective ex vivo storage method for a wide variety of solid organs from various species.8–11 However, the tolerance of feline kidneys to CS is unknown, although in 1 case series,12 postoperative graft function was described as excellent for kidneys that had been preserved by CS for up to 7 hours prior to transplantation. In dogs under ideal conditions, successful graft function has been achieved following transplantation of kidneys stored for up to 6 days in simple static CS and up to 7 days with the use of continuous perfusion methods.13–15
In current clinical practice and most experimental studies, long-term storage of feline kidneys intended for transplantation is not necessary because the ex vivo period for most grafts is measured in minutes rather than hours or days. Therefore, the magnitude of any potential advantage provided by CS on the immediate function of a graft following transplantation is unknown. The sensitivity of feline kidneys to CS has not been established. Additionally, in cats, the effect of short-term warm ischemia on graft function, compared with the effects of confounders associated with the surgical procedure such as ureteral spasm and transient ureteral obstruction, is likewise unknown. Ultrasonographic evaluation of renal grafts is a rapid and sensitive method that can be used to facilitate assessment of those confounding factors on graft function following transplantation.
The RI is a unitless measure that describes the pulsatility of the intrarenal arterial time-velocity waveform and is independent of both insonation angle and Doppler frequency.16 Sonography and measurement of the RI have been evaluated in healthy cats,17 cats with renal disease,18 and cats following renal autotransplantation19 and clinical renal transplantation.20 The RI is most useful for the identification of azotemic cats with nonobstructive renal disease and dogs with acute tubular necrosis,18 and is considered a useful adjunct diagnostic modality for assessing renal function in human patients following kidney transplantation21 and those with chronic nephropathies.22 In cats that underwent kidney transplantation, graft size but not RI increased significantly after autotransplantation16,19 and for allografts in recipients with clinical signs of graft rejection or ureteral obstruction.20 Because both RI and kidney size are correlated with renal disease in cats, those may be 2 additional useful measurements for prediction of short-term outcomes for cats that receive grafts by the IT procedure or following CS.
The purpose of the study reported here was to describe the use of simple CS methods for short periods on immediate posttransplantation function of renal autografts in cats. We hypothesized that graft function would be significantly better in cats receiving kidneys that underwent CS in a sucrose phosphate preservation solution for periods that mimicked most clinical or experimental scenarios than in cats receiving kidneys during an IT procedure.
Materials and Methods
Animals
All study procedures were reviewed and approved by the University of Wisconsin Animal Care and Use Committee. Fifteen male domestic shorthair cats were used in the study. All cats were approximately 1 year old, weighed between 3.0 and 4.0 kg, and were considered healthy on the basis of results of a physical examination and serum biochemical analysis.
Experimental design
The surgical model used consisted of autotransplantation of the left kidney and nephrectomy of the right kidney. All surgeries were performed by 1 surgeon (JFM). Each cat was randomly assigned to either the IT or CS group by use of a random-number generator. For cats in the IT group (n = 6), the left kidney was removed, the end of the renal artery was cleared of blood by brief topical lavage with saline (0.9% NaCl) solution, and the vascular anastomosis was begun. For cats in the CS group (n = 9), the left kidney was removed, flushed with a sucrose phosphate preservation solution (preservation solution) as described,23 and then stored in the preservation solution on ice while the vascular anastomosis site was prepared.
Anesthesia and supportive care
All cats were premedicated with acepromazine maleate (0.02 mg/kg, IM) and butorphanol tartratea (0.2 mg/kg, IM). Anesthesia was induced with thiopental sodium (10 mg/kg, IV) and maintained with isoflurane in oxygen delivered via an endotracheal tube. Cefazolin sodium (22 mg/kg, IV) was administered 30 minutes before surgery. Mannitol (0.5 g/kg, IV) was administered 10 minutes before removal of the left kidney and again just prior to release of the vascular clamps at completion of the vascular anastomosis. A continuous IV infusion of lactated Ringer solution (5 mL/kg/h) was administered during surgery; the infusion was slowed to 2 to 4 mL/kg/h for 12 to 18 hours after surgery and then provided by SC administration thereafter as needed. Analgesia was provided by application of a transdermal fentanyl patch (25 μg/h) prior to surgery and supplemented with butorphanola (0.2 mg/kg, IM, q 4 to 6 h) after surgery as needed.
Surgical techniques
For each cat, a ventral midline celiotomy was performed, and the kidneys, ureters, urinary bladder, and other internal organs were examined. The left kidney was harvested for autotransplantation. In preparation for harvest, the left kidney, renal vessels, and ureter were isolated from all peritoneal attachments. The ureter was transected distally at the level of the bladder, and the remaining stump was double ligated with 4–0 silk. The kidney was not removed (harvested) until the blood pressure was within reference limits and urine was visible from the transected ureter.
For cats in the IT group, the implantation site was prepared after the kidney, renal vessels, and ureter had been isolated but before the renal vessels were clamped and divided to minimize the duration of warm ischemia as much as possible. Implant site preparation included excising excess adventitia from the vena cava and aortic walls, partially occluding the vena cava with an atraumatic vascular clamp, fenestrating the caval wall, and preplacing vascular sutures for attachment of the vein.
Implantation was performed on the left side with end-to-side anastomosis of the renal artery and vein to the aorta and vena cava, respectively. Arterial and venous anastomoses were performed by use of a continuous suture pattern with 8–0 nylon and 10–0 braided polyester suture material, respectively. The time required for vascular anastomosis was calculated as the duration between excision of the left kidney and removal of vascular clamps following completion of anastomoses for the IT group and as the duration between removal of the kidney from the preservation solution and removal of vascular clamps following completion of anastomoses for the CS group. For all cats, the renal vein was anastomosed to the vena cava first, then a completely occluding vascular clamp was placed on the aorta and the renal artery was anastomosed to the aorta. Neoureterocystostomy was performed by use of an intravesicular technique. The bladder wall was incised, and the ureter from the graft was sutured to the bladder mucosa in a simple continuous pattern with 8–0 polyglactin 910.b The bladder wall was then closed in 2 layers by use of 5–0 polyglactin 910 in a simple continuous pattern. The graft was stabilized by suturing the renal capsule to the retroperitoneal membrane by use of 5–0 polyglactin 910 in a simple interrupted pattern. After implantation of the graft, the contralateral (right) kidney was excised by use of standard surgical techniques. The graft anastomoses were rechecked for hemorrhage or other problems, and the abdominal cavity was closed in a routine manner.
Graft preservation protocol for the CS group
The preservation solution (Appendix) was prepared as described23 and sterilized by filtration (filter pore size, 0.22 μm) into a sterile plastic IV bag and stored refrigerated at 5°C. A separate bag of preservation solution was prepared for each cat in the CS group. All prepared solutions were analyzed prior to use and found to have a pH within ± 0.01 and osmolarity within ± 3 mOsmol/L of the desired levels.
Immediately after the left kidney was removed from the abdominal cavity, the left renal artery was cannulated with an 18-gauge Teflon catheter and flushed from a height of 100 cm (pressure, approx 100 cm H2O) with 15 to 30 mL of cold (approx 10°C) preservation fluid. Flushing was stopped as soon as the venous effluent was visibly clear of blood or the kidney developed excess turgor as determined by palpation, which was suggestive of the formation of flush-induced edema. The kidney was then immersed in preservation solution in a stainless steel bowl nestled inside a larger stainless steel bowl that contained a sterile ice and saline slurry. The bowl containing the kidney and preservation solution was agitated by manual swirling until the preservation solution became palpably colder, indicating that the temperature of the preservation solution had equilibrated with that (approx 0 to 2°C) of the ice slurry by heat transfer through the wall of the steel bowl. The kidney was not flushed with any additional preservation solution prior to implantation. Before implantation, the renal vessels were prepared by excising excess adventitia and squaring of the vessel ends while the kidney remained submerged in the cold preservation solution. The graft was then implanted in the same manner as that described for the IT group.
Postoperative monitoring and data collection
For all cats, postoperative monitoring included a physical examination and determination of PCV, and blood total solids, BUN, and serum creatinine concentrations on a daily basis for 14 days. The daily urine output was subjectively measured as an indication of ureteral patency.
The autograft was ultrasonographically examined at 1, 2, 4, 7, 9, and 11 days after surgery or as needed for cats with abnormally increased BUN or creatinine concentrations to ascertain whether those abnormalities were related to surgical complications or other technical problems. A registered diagnostic medical sonographer (FAD), who was unaware of (blinded to) the treatment group assignment and clinical and biochemical measures for each cat, performed all ultrasonographic examinations. Cats were positioned in dorsal recumbency in a trough for each examination. The same ultrasound machinec was used for all examinations. A curvilinear transducerd was used at a 7-MHz center frequency for all imaging except assessment of the corticomedullary distinction, which was evaluated with a linear array transducere at 11 MHz. All machine settings (frequency, overall gain, segmental gain, depth, dynamic range, and all power Doppler settings) were standardized and held constant throughout the study. Resistive index values were calculated by use of an arterial waveform obtained with duplex Doppler imaging of an arcuate artery in accordance with the following standard equation: RI = (maximum arterial blood velocity – minimum arterial blood velocity)/maximum arterial blood velocity. Resistive index was calculated by the ultrasound machine software at the time of each examination. At least 2 RI measurements were performed for different arteries during each examination. The mean of all RI measurements obtained during an examination was calculated and used as the RI for that examination.
Graft length was defined as the length from the cranial to caudal pole of the kidney on a midsagittal image. Cross-sectional area was defined as the product of two orthogonal measurements of graft width obtained from a transverse image at the level of the renal pelvis. Graft volume was defined as the product of the graft length, cross-sectional area, and a constant (0.5236) used to approximate the volume of a prolate spheroid.24 The size of the renal pelvis was also determined.
Cats that developed progressive uremia that was unresponsive to basic supportive measures were euthanized, and all cats that were still alive at 2 weeks after surgery were likewise euthanized. Euthanasia was performed by administration of a commercial euthanasia solution (10 mL/kg, IV).
Data analysis
Peak and mean serum creatinine and BUN concentrations were determined for each treatment group each day after surgery. Longitudinal and in-tergroup differences in serum creatinine and BUN concentrations, postoperative RI, and graft length, cross-sectional area, and volume were assessed by repeated-measures ANOVA. Values of P ≤ 0.05 were considered significant for all analyses.
Results
Six cats (2 in the IT group and 4 in the CS group) developed complications that precluded evaluation of immediate graft function and were excluded from all analyses. Those complications included ureteral problems such as urine leakage (n = 1 cat in the CS group) and ureteral obstruction (2 cats in the CS group), arterial thromboses and graft infarction (2 cats in the IT group), and substantial postoperative hemorrhage followed by central vein thromboembolism 4 days after surgery (1 cat in the CS group). Thus, for analysis purposes, there were 4 and 5 cats in the IT and CS groups, respectively.
The mean ± SE time required for vascular anastomosis for the IT group (54.0 ± 2.7 minutes) did not differ significantly from that for the CS group (53.7 ± 3.7 minutes). For the CS group, the mean ± SE duration of kidney storage was 24.4 ± 4.8 minutes.
The mean ± SE serum creatinine concentration for the cats in the IT group was significantly greater than that for the cats in the CS group for the duration of the postoperative observation period (Figure 1). The serum creatinine concentration typically peaked 2 or 3 days after surgery for cats in the IT group and between days 1 and 3 after surgery for cats in the CS group. The mean ± SE peak serum creatinine concentration for the IT group (10.7 ± 2.9 mg/dL) occurred 3 days after surgery, whereas that for the CS group (2.58 ± 0.44 mg/dL) occurred 1 day after surgery. The mean serum creatinine concentration for both groups decreased from the respective peak concentrations throughout the remainder of the observation period. On day 14 after surgery (immediately before euthanasia), the mean ± SE serum creatinine concentration for the IT group (2.32 ± 0.11 mg/dL) remained increased from the laboratory reference range (0.7 to 1.9 mg/dL), whereas that for the CS group (1.45 ± 0.14 mg/dL) was within the reference range.
Mean ± SE postoperative serum creatinine (A) and BUN (B) concentrations over time for 4 of 6 cats that underwent autotransplantation of the left kidney by use of an IT procedure (IT group; black circles) and 5 of 9 cats that underwent autotransplantation of the left kidney after it had undergone a period of CS (CS group; white squares). All cats were healthy prior to surgery and underwent nephrectomy of the right kidney during the same procedure in which the left kidney was autotransplanted. Six cats (2 in the IT group and 4 in the CS group) were excluded from the analyses because they developed complications that precluded evaluation of immediate graft function such as urine leakage from the ureter (n = 1 cat in the CS group), ureteral obstruction (2 cats in the CS group), arterial thromboses and graft infarction (2 cats in the IT group), and substantial postoperative hemorrhage followed by central vein thromboembolism (1 cat in the CS group). *Within a day, the respective values for the 2 groups differ significantly (P ≤ 0.05).
Citation: American Journal of Veterinary Research 78, 3; 10.2460/ajvr.78.3.330
The postoperative BUN concentrations were more variable than the postoperative serum creatinine concentrations. The mean ± SE BUN concentration for the IT group was significantly greater than that for the CS group on days 1 through 5 and again on days 10 through 14 after surgery (Figure 1). The BUN concentration peaked between days 1 and 6 after surgery for cats in the IT group and on day 2 for cats in the CS group. The mean ± SE peak BUN concentration for the IT group (115 ± 37 mg/dL) occurred 4 days after surgery, whereas that for the CS group (37 ± 3.2 mg/dL) occurred 1 day after surgery. The mean BUN concentration for both groups decreased from the respective peak concentrations throughout the remainder of the observation period. On day 14 after surgery, the BUN concentration was within the laboratory reference range (20 to 30 mg/dL) for all 5 cats in the CS group but only 2 of 4 cats in the IT group.
Variables derived from data obtained during postoperative ultrasonographic examinations were summarized (Table 1). Five cats in the IT group and 7 cats in the CS group underwent at least 1 ultrasonographic examination of the graft after transplantation; however, all 6 scheduled posttransplantation ultrasonographic examinations were performed on only 4 cats in the IT group and 3 cats in the CS group because of the limited availability of the ultrasound machine. Only data from those 7 cats with 6 complete posttransplantation ultrasonographic examinations were analyzed and reported. The renal pelvis of the graft was intermittently measured for 3 of the 4 cats in the IT group and all 3 cats in the CS group. When the renal pelvis data were aggregated, the mean ± SE size of the renal pelvis for the IT group (3.2 ± 0.26 mm) did not differ significantly (P = 0.89) from that for the CS group (2.9 ± 0.33 mm). The RI values did not change over time. Graft cross-sectional area and volume varied significantly during the observation period for the IT group but not the CS group. The mean RI for the CS group was significantly greater than that for the IT group 11 days after transplantation. The mean graft length for the CS group was significantly shorter than that for the IT group 2 days after transplantation. The mean graft cross-sectional area for the CS group was significantly smaller than that for the IT group 11 days after surgery. Similarly, the mean graft volume for the CS group was significantly less than that for the IT group at 2 and 11 days after surgery.
Descriptive data for renal autografts on days 1, 2, 4, 7, 9, and 11 after transplantation for 4 of 6 cats that underwent an IT procedure (IT group) and 3 of 9 cats in which the graft was transplanted after a period of CS (CS group).
| Group | ||||
|---|---|---|---|---|
| Variable | Day after transplantation | IT | CS | All cats |
| RI | 1 | 0.586 ± 0.00 | 0.583 ± 0.11 | 0.584 ± 0.06 |
| 2 | 0.590 ± 0.01 | 0.472 ± 0.05 | 0.539 ± 0.03 | |
| 4 | 0.527 ± 0.02 | 0.440 ± 0.06 | 0.484 ± 0.03 | |
| 7 | 0.561 ± 0.02 | 0.478 ± 0.08 | 0.525 ± 0.04 | |
| 9 | 0.568 ± 0.05 | 0.538 ± 0.06 | 0.553 ± 0.03 | |
| 11 | 0.553 ± 0.04 | 0.584 ± 0.05* | 0.566 ± 0.03 | |
| Graft length (mm) | 1 | 41.90 ± 2.5 | 40.73 ± 1.1 | 41.32 ± 1.3 |
| 2 | 44.78 ± 2.4 | 41.97 ± 2.2* | 43.57 ± 1.6 | |
| 4 | 45.48 ± 2.1 | 43.03 ± 3.0 | 44.43 ± 1.7 | |
| 7 | 44.53 ± 2.3 | 43.13 ± 1.5 | 43.93 ± 1.4 | |
| 9 | 45.08 ± 2.9 | 40.83 ± 0.92 | 43.26 ± 1.8 | |
| 11 | 43.30 ± 2.6 | 40.90 ± 2.6 | 42.27 ± 1.8 | |
| Graft cross-sectional | 1 | 848.2 ± 147.6a | 813.3 ± 167.5 | 830.7 ± 100.0 |
| area (mm2) | 2 | 985.6 ± 116.2a,b | 893.4 ± 118.9 | 946.1 ± 78.9 |
| 4 | 1,082 ± 125.2b | 880.6 ± 169.7 | 995.8 ± 101.2 | |
| 7 | 972.1 ± 143.9a,b | 771.5 ± 135.6 | 886.1 ± 99.1 | |
| 9 | 916.8 ± 160.0a | 750.0 ± 93.5 | 845.3 ± 98.5 | |
| 11 | 938.5 ± 133.3a,b | 777.9 ± 105.6* | 869.6 ± 87.9 | |
| Graft volume (mm3) | 1 | 18,988 ± 4,153a,b | 17,528 ± 3,938 | 18,258 ± 2,580a,b |
| 2 | 23,538 ± 3,843a,b | 19,909 ± 3,526* | 21,983 ± 2,556a,b | |
| 4 | 26,151 ± 3,977b | 20,334 ± 5,122 | 23,658 ± 3,106b | |
| 7 | 23,178 ± 4,511a,b | 17,609 ± 3,384 | 20,791 ± 2,952a,b | |
| 9 | 22,346 ± 5,082a,b | 16,125 ± 2,362 | 19,680 ± 3,123a | |
| 11 | 21,792 ± 4,081a | 16,941 ± 3,913* | 19,713 ± 2,679a | |
Values represent the mean ± SE. All cats were healthy prior to surgery and underwent autotransplantation of the left kidney and nephrectomy of the right kidney. Only data for cats that underwent all 6 scheduled posttransplantation ultrasonographic examinations were analyzed. All ultrasonographic examinations were performed by a registered diagnostic medical sonographer who was unaware of the treatment group assignment for each cat, and all machine settings (frequency, overall gain, segmental gain, depth, dynamic range, and all power Doppler settings) were standardized and held constant throughout the study. Resistive index values were calculated by use of an arterial waveform obtained with duplex Doppler imaging of an arcuate artery in accordance with the following standard equation: RI = (maximum arterial blood velocity – minimum arterial blood velocity)/maximum arterial blood velocity. At least 2 RI measurements were performed for different arteries during each examination, and the mean of all RI measurements obtained during an examination was calculated and used as the RI for that examination. Graft length was defined as the length from the cranial to caudal pole of the kidney on a midsagittal image. Cross-sectional area was defined as the product of 2 orthogonal measurements of graft width obtained from a transverse image at the level of the renal pelvis. Graft volume was defined as the product of the graft length, cross-sectional area, and a constant (0.5236) used to approximate the volume of a prolate spheroid.
Value differs significantly (P ≤ 0.05) from the corresponding value for the IT group.
Within a group and variable, values with different superscripts differ significantly (P ≤ 0.05); the absence of superscript letters indicates that the values did not differ significantly over time for that variable within that group.
Discussion
To our knowledge, the present study was the first controlled study involving cats to evaluate the effect of short-term CS of renal autografts on immediate posttransplantation graft function. Results indicated that immediate graft function for kidneys that underwent short-term CS was markedly better than that for kidneys that underwent IT and a period of warm ischemia.
The sucrose phosphate preservation solution with which the kidneys were flushed and immersed immediately after harvest allowed the grafts to be stored at a low temperature during the fairly short ex vivo period without significantly impairing graft function immediately after transplantation. The apparent protective effect of CS of grafts in the preservation solution has the potential to improve both the short- and long-term outcomes for feline renal transplant patients, as well as simplifying the logistics associated with the performance of the procedure. This was the first study to provide a protocol for CS preservation of renal autografts and validate the benefits of such preservation on graft function immediately following autotransplantation by use of serial serum biochemical analyses and ultrasonographic examinations.
Short-term CS of renal grafts as part of either a clinical or experimental renal transplantation protocol in cats has a number of advantages in addition to protection of the grafts from the adverse effects induced by warm ischemia. It allows the harvest of the donor kidney and preparation of the recipient for graft implantation to be performed serially rather than in parallel, which decreases the demand for operating rooms, hospital equipment, and skilled personnel and allows the surgical team to take a break between donor and recipient procedures. Cold storage also allows the graft vessels to be prepared (ends trimmed and loose adventitia removed) either in the preservation solution or on a gauze sponge before being reimmersed in the cold preservation solution and preparation of the recipient vessels without the pressure of trying to minimize the time required for vascular anastomoses so as to prevent or minimize ischemia-induced adverse effects on graft function and survivability. Additionally, our laboratory group has developed and described novel modifications to the neoureterocystotomy procedure so that the ureteral papilla implantation can be easily performed during CS.25 The only obvious disadvantages to CS are the need to prepare the preservation solution and implement an additional step (graft flushing and storage) within the complex transplantation protocol.
In the present study, the immediate posttransplantation graft function was better for kidneys that underwent CS, compared with that for kidneys that underwent IT, presumably because the grafts in CS did not incur warm ischemic injury during the nearly 1 hour required for vascular anastomoses. Kidneys that undergo periods of warm ischemia generally develop acute tubular necrosis, which was likely the cause of impaired function for many of the grafts in this study. Interestingly, the renal autografts in the CS group were significantly smaller than the autografts in the IT group, even though the grafts in the IT group did not have ultrasonographic evidence of hydronephrosis. In cats, the cross-sectional area of renal allografts increases during acute rejection of the allograft.26 The fact that the graft cross-sectional area and volume for the cats in the IT group were significantly greater than those for the cats in CS group 11 days after transplantation might suggest that the extent of parenchymal inflammation or edema was greater for grafts in the IT group than for grafts in the CS group. Although it is unknown whether increasing graft size is associated with retroperitoneal fibrosis observed in some cats following renal transplantation, it may help trigger graft rejection.26
The renal autografts of the present study were not retrieved for histologic examination until the serum creatinine and BUN concentrations had returned to within or nearly within the respective reference ranges. Consequently, the only histologic abnormalities detected were those associated with nonspecific inflammatory changes and loss of renal tubules (data not presented). Specific histologic changes caused by acute graft dysfunction immediately after transplantation could not be ascertained. The peak concentrations of both serum creatinine and BUN for the cats in the IT group were approximately 5 times those for the cats in the CS group. It is possible that warm ischemia induced in the grafts during the transplantation procedure for the IT group resulted in a variety of effects that collectively caused such substantial increases in the biochemical indices of renal function. For example, in addition to tubular injury, warm ischemia may promote ureteral spasm, which can obstruct urine outflow from the graft. Results of another study5 suggest that serum creatinine concentrations of cats following renal transplantation were positively associated with both ureteral spasm and warm ischemic injury. Regardless, the results of the present study indicated the posttransplantation serum creatinine and BUN concentrations for cats that received autografts that had undergone short-term CS were increased only slightly from the respective reference ranges for a short period of time, whereas those for cats that received autografts that underwent a period of warm ischemia during an IT procedure were substantially increased from the respective reference ranges for a fairly long period (up to 1 week), which suggested that CS helped to preserve graft function.
In a clinical setting, some degree of acute graft injury is tolerable in cats that have an otherwise uncomplicated recovery from a renal transplant procedure (as evidenced by the multiple successful renal transplants that have been performed in cats without the use of CS), and a full recovery of graft function over time is usually expected in such patients. Unfortunately, many cats develop unstable cardiovascular function after receiving a renal transplant and undergo hypertensive or hypotensive episodes,27 which adversely affect graft perfusion and may exacerbate any warm ischemic injury. In unstable patients, the likelihood of postoperative graft dysfunction is positively associated with the extent of graft injury induced by warm ischemia during the transplant procedure. Therefore, CS of grafts, even for a short period, may provide a safety buffer for maintenance of graft function in cats receiving renal transplants.
It should be noted that cold temperature can adversely affect cell survival.28 The detrimental effects of cold on cell survival have been described in experimental models for various organ and vascular grafts, particularly when the grafts are flushed with and stored in physiologic saline solution.29 Consequently, specialized preservation solutions have been developed that minimize the adverse effects associated with cold ex vivo storage.
The basic principles of cold preservation involve addressing or minimizing the 3 primary mechanisms of cold injury. Preservation solutions must be formulated to suppress cell swelling induced by cold temperatures. This is most commonly done by replacing chloride with an impermeable molecule such as gluconate or lactobionate as the major anion in a solution or by use of an osmotic agent, such as sucrose,30,31 as was done in the present study. Preservation solutions must also provide metabolic support; providing for energy metabolism is particularly important for tissues intended for long-term storage.32–34 Finally, biological manipulation of tissues through cell signaling pathways during ex vivo storage has been recognized as a key factor for optimization of preservation protocols but is likely not necessary for tissues that will undergo only short-term CS such as those used in most veterinary transplant procedures.13 Other mechanisms of tissue injury associated with CS include derangements in intracellular calcium metabolism, oxidative stress, and mitochondrial function35–38; however, the extent to which those mechanisms contribute to the injury of grafts stored for short periods is unclear.
Each organ has a finite tolerance to currently used CS methods, with the heart having the shortest tolerance and the kidney having the longest tolerance.13,14,39 The tolerance of specific organs to cold-induced injury also varies substantially among species.40 Among species commonly used for experimental study of organ storage or clinical organ transplantation, rat organs appear to be the most susceptible to CS-induced injury followed by porcine organs, whereas canine organs appear to be the least susceptible to CS-induced injury.40 The tolerance of human organs to CS appears to be intermediate between that of porcine and canine organs.40 The feasible duration that feline kidneys or other organs can be stored is unknown, but results of the present study suggested that feline kidneys tolerated CS well for up to 24 minutes as evidenced by the minimal effects on renal function observed immediately following transplantation. Canine kidneys have been successfully stored at low temperatures for up to 7 days by use of simple flushing techniques or continuous arterial perfusion with fairly complex preservation solutions.13–15,35 If similar CS times can be achieved for feline kidneys, a kidney donation program in which kidneys are harvested from cats that are euthanized for reasons other than renal disease (ie, analogous to organ procurement programs in human medicine) might become feasible on a limited basis. However, aside from the technical aspects of organ preservation, the logistics of such a program are likely to make that approach impractical for most veterinary facilities.
The extent of warm ischemic injury incurred by feline renal grafts appears to be dependent on the duration of ischemia. Although we recognize the limitations of comparing results between studies, we plotted peak serum creatinine concentration versus time required for vascular anastomosis for the cats of both the IT and CS groups of the present study and the cats of another study5 that underwent the same renal autotransplantation procedure used for the IT group (Figure 2). We used peak serum creatinine concentration as a surrogate for extent of graft injury. The serum creatinine concentration (and presumably extent of graft injury) increased as the vascular anastomosis time increased, and the magnitude of the increase in creatinine concentration for the cats that underwent the IT procedure was substantially greater than that for cats in the CS group. Those findings indicated the importance of minimizing the time required for vascular anastomosis in renal transplantation procedures and provided further incentive for the use of technologies that decrease graft injury and extend the duration of ex vivo graft survival.
Mean preoperative (0 minutes) and peak serum creatinine concentration versus vascular anastomosis time for the cats of Figure 1 (black circles) and 6 cats of another study5 that underwent the same renal autotransplantation procedure used for the IT group of the present study (white square). The dashed line represents the regression line for the CS group, and the solid line represents the regression line for the cats that underwent the IT procedure in both studies. Notice that the serum creatinine concentration (and presumably extent of graft injury) increased as the vascular anastomosis time increased and that the magnitude of that increase for the cats that underwent the IT procedure was substantially greater than that for cats in the CS group.
Citation: American Journal of Veterinary Research 78, 3; 10.2460/ajvr.78.3.330
The severity of graft injury represents the cumulative effects of all stressors incurred by the graft during the perioperative period including harvest and implantation. Clearly, graft injury caused by ischemia is time dependent, and the rate of cumulative injury slows as temperature decreases. In the field of transplantation medicine, terms have been developed to describe the various ischemic stressors incurred by grafts such as warm ischemia time, cold ischemia time, and second warm ischemia time. Warm ischemia time is generally only a few minutes for grafts that undergo some type of CS but is much longer for grafts that do not undergo CS because, for those grafts, it includes the vascular anastomosis time and any time required for site preparation. Cold ischemia time is the duration of ex vivo CS. Second warm ischemia time is the period during which a graft that underwent CS slowly warms up because of exposure to room temperature while it is being implanted and is generally equivalent to the vascular anastomosis time. For grafts that undergo CS, the extent of injury incurred during the second warm ischemia time is much less than that incurred during the warm ischemia time for grafts that do not undergo CS. This is because the rate of graft injury is positively associated with temperature. The temperature of grafts coming out of CS during the second warm ischemia time slowly increases to room temperature until reperfusion is established, whereas the temperature of grafts that do not undergo CS remains at or close to body temperature, which is typically much higher than room temperature.
In addition to improved immediate graft function, protection of renal grafts from warm ischemic injury may have long-range implications on graft survival. In human transplant patients, delayed graft function following transplantation is positively associated with the rate of acute rejection episodes and the likelihood of chronic rejection, which is characterized by graft vascular disease.41–45 The incidence of graft vascular disease in feline transplant patients is unknown. Additional studies are necessary to determine the rate of chronic graft disease in cats, and identifying methods to reduce it will be important for improving long-term patient management.
The overall rate of surgical complications in the cats of the present study was high (6/15 [40%]). This was a reflection of the challenging nature of the renal transplantation procedure and the learning curve associated with the ureteral implantation techniques for the study personnel. Surgical complications inherent to the renal transplantation procedure can also affect immediate graft function. Therefore, determining whether immediate graft dysfunction is the result of ischemia-induced parenchymal injury or technical or procedural problems can be difficult clinically, as evidenced by similarly high complication rates in experimental studies4–6 in which various models that were developed to differentiate the effects of graft ischemia from the effects of ureteral complications were assessed.
The preservation solution used for the CS procedure of the present study was inexpensive and easily compounded from ingredients that are readily available from any commercial laboratory chemical supplier. It took very little time and required no specialized skills or equipment to prepare. The preservation solution was usually prepared the day before the transplant procedure and placed in a refrigerator so that it would be cooled for use. Thus, preparation and use of this preservation solution should be practical and readily applicable in any clinical or laboratory setting.
Limitations of the present study included the small number of cats enrolled in the study and the relatively high proportion of cats that developed complications, which prevented evaluation of immediate graft function and caused them to be excluded from the analysis. Also, all the cats of the present study were young and healthy, and the magnitude of the results may not be indicative of those for clinical patients, which typically have substantial comorbidities and frequently develop complications that can affect serum biochemical variables and hinder assessment of graft function.
Results of the present study indicated that, in cats, immediate posttransplantation function of renal autografts that underwent CS, even for a short period of time, was significantly better than that of renal autografts that were IT and did not undergo CS. Improved graft function immediately after transplantation should translate into an increased likelihood of a successful long-term outcome for transplant patients. This study also provided baseline performance data that can be used for comparison purposes in the future. Additionally, cats have a limited number of blood types and metabolize immunosuppressive drugs in a manner similar to humans; therefore, renal transplantation in cats may serve as a desirable model for investigating the long-term effects of renal transplantation in human patients.
Acknowledgments
Supported by the University of Wisconsin Companion Animal Fund.
The authors declare that there were no conflicts of interest. Presented as a poster presentation at the American College of Veterinary Surgeons Symposium, Denver, October 2004.
ABBREVIATIONS
| CS | Cold storage |
| IT | Immediate transplantation |
| RI | Resistive index |
Footnotes
Torbugesic, Fort Dodge Animal Health, Fort Dodge, Iowa.
Vicryl, Ethicon Inc, Somerville, NJ.
GE Logiq 400, General Electric Co, Piscataway, NJ.
C721, General Electric Co, Piscataway, NJ.
LA39, General Electric Co, Piscataway, NJ.
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Appendix
Composition of the preservation solution that was used to flush feline renal autografts immediately after harvest and in which the autografts were immersed during CS.
| Component | Concentration |
|---|---|
| NaH2PO4 (monobasic) | 15.5 mmol/L |
| Na2HPO4 (dibasic) | 53.6 mmol/L |
| Sucrose | 140.0 mmol/L |
| Heparin | 1,000 U/L |
| Distilled water | — |
— = Not applicable.
Compounds were mixed and allowed to equilibrate for a short period. The volume prepared was 500 mL/preparation. The pH was adjusted to 7.2 as necessary by the addition of NaOH or HCl in a dropwise manner. Then, the solution was sterilized by filtration (filter pore size, 0.22 μm) into a sterile plastic IV bag and stored refrigerated at 5°C. A separate bag of preservation solution was prepared for each cat in the CS group.
