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Schaub, Meike1 3; Ploetz, Christian J.1; Gerbaulet, Daniel1; Fang, Liu1; Kranich, Pia2; Stadlbauer, Thomas H. W.2; Goettman, Uwe1; Yard, Benito A.1; Braun, Claude1; Schnuelle, Peter1; van der Woude, Fokko J.1

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doi: 10.1097/01.TP.0000119164.47302.49
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Survival of kidneys from brain-dead donors is markedly inferior to survival of kidneys from living donors (1). Therefore, brain death is regarded as an independent risk factor for chronic allograft dysfunction. These findings have been extended by different groups studying the influence of brain death on chronic rejection in animal models (2–4). The underlying concept is that brain death induces a “state of inflammation” in several tissues, which leads to an increased immunogenicity of a potential graft. Therefore, in addition to improved immunosuppression of the recipient, preconditioning of the graft could become an important therapeutic strategy to improve long-term graft acceptance.

In two retrospective clinical studies, Schnülle et al. (5, 6) demonstrated that dopamine treatment of the brain-dead donor results in better long-term renal allograft survival. There are conflicting data in the literature regarding the mode of action and effects of dopamine in clinical kidney transplantation (7–9). In addition to its hemodynamic properties, dopamine is known to indirectly exert an antioxidative effect. During its metabolism, dopamine is degraded to quinones and semi-quinones, leading to the production of reactive oxygen species such as H2O2 and O2- (10). These reactive oxygen species in turn result in the induction of endogenous antioxidatives such as heme oxygenase (HO)-1 (11). HO is an enzyme responsible for degradation of heme. Its inducible isoform HO-1 is induced in tissues by various stimuli, many of which are pro-oxidant. In recent years it has been described that induction of HO-1 exerts protective effects in various forms of tissue injury in different animal models (12).

Berger et al. (13) demonstrated that incubation of human umbilical vein endothelial cells with dopamine induces HO-1. In different in vitro studies it was also shown that dopamine down-regulates chemokine production under proinflammatory conditions (14). The mechanisms of the protective effects of dopamine treatment have not been studied in vivo. To further elucidate the mechanisms by which dopamine exerts its effects on kidneys of brain-dead donors in vivo, we set up a model for brain death in rats. Brain death was induced by epidural inflation of a 3F Fogarty catheter. Apneic animals were mechanically ventilated for 6 hr, and hemodynamics, blood gases, and hematocrit levels were monitored. Ventilated, non–brain-dead animals served as controls. According to clinical application, dopamine was given in different dosages (2, 6, 10, or 14 μg/kg/min).

We tested the hypothesis that clinically relevant dosages of dopamine reduce the immunogenicity and enhance the anti-oxidative capacity of kidneys of brain-dead donors.


Animal Studies

Experiments were performed in male Fisher 344 rats weighing 200 to 250 g. Animals were purchased from Harlan-Winkelmann GmbH, Borchen, Germany. Studies were performed according to the “Guide for the Care and Use of Laboratory Animals,” published by the National Institutes of Health (publication number 85–23, revised 1996), and were approved by German federal regulations (RP Karlsruhe, AZ 35–9185.81/60/90).

Before the induction of brain death, animals were anesthetized with ketamine (Ketanest, Pfizer, Karlsruhe, Germany, 100 mg/kg intraperitoneally) and xylazine (Rompun, BayerVital, Leverkusen, Germany, 6 mg/kg intraperitoneally) and placed on a heating table to keep their body temperature constant. In an occipital burr hole, a 3F Fogarty catheter was inserted epidurally and inflated with 200 μL of saline. While cardiocirculatory functions were preserved, animals became apneic and were mechanically ventilated by a tracheostoma with a rodent ventilator (Ugo Basile, Comerio, Italy). Adequate ventilation was monitored by blood gas analysis during the experiment. Brain death was verified by electroencephalogram monitoring (EEG Coupler, Alfos AG, Biel-Benken, Germany) as cessation of neural activity. Systemic blood pressure (mean arterial pressure [MAP], mm HG) was continuously measured by an inguinal arterial catheter (Statham pressure transducer P23Db and a Gould pressure processor, FMI, Ober-Beerbach, Germany). Renal blood flow (RBF, milliliters/minute) was monitored by an ultrasonic transit-time flow probe (1RB, Transonic Systems Inc., Ithaca, NY) placed around the left renal artery. Animals were ventilated and monitored for 6 hr. Insufficient ventilation or a decrease in hemoglobin level of greater than 2 mg/dL led to exclusion from the experiment. Deeply anesthetized, apneic ventilated animals without induction of brain death served as controls.

Dopamine was given through a catheter in the femoral vein in four different dosages (2, 6, 10, and 14 μg/kg body weight/min). After 6 hr, only the nonmanipulated right kidney was explanted and snap-frozen in liquid nitrogen for in vitro analysis. Each group (brain-dead animals, ventilated animals, ventilated animals with dopamine 10 μg/kg/min, and four groups of brain dead animals receiving different dosages of dopamine mentioned above) consisted of a minimum of five animals, and some groups had up to seven animals.


Serial cryostat sections (5 μm) were fixed in 95% ethanol for immunohistochemical staining. After phosphate-buffered saline rinsing and blockade of endogenous peroxidase (3% hydrogen peroxide) and endogenous biotin (Avidin blocking kit, Vector, Burlingame, CA), sections were incubated for 2 hr with primary antibody for monocytes and macrophages ED1 (monoclonal mouse anti-rat, Serotec, Eching, Germany), major histocompatibility complex (MHC) II (F-17–23–2, monoclonal mouse anti-rat, Serotec), and P-selectin (PharMingen, Heidelberg, Germany), followed by incubation with species-specific secondary antibodies for 1 hr. Irrelevant murine or rabbit control antibody or omission of the primary antibody served as controls. Standard avidin-biotin complex staining was performed according to the manufacturer’s instructions (ABC kit, Vector); 3,3′ diaminobenzidine substrate was applied, and the sections were counterstained with hematoxylin. After dehydration in graded ethanol and xylene, slides were mounted in Permount for evaluation. ED1 and MHC II positive cells were counted in a blinded fashion under the microscope at 400× enlargement. At least five animals per group and 15 fields or more per view per sample were evaluated in a blinded fashion. P-selectin expression in renal vessels was graded from 0 (no staining) to 3+ (strong staining) by blinded investigators, and the results were expressed as the percentage of total numbers per group per score.

Western Blot Analysis

Frozen tissue samples were covered with a protease inhibitor (Complete Mini, Roche, Mannheim, Germany) and homogenized with a tissue homogenizer (Polytron, IKA Labortechnik/Fischer Scientific, Schwerte, Germany).

Protein concentration was determined according to Bradford (Protein Assay, Bio-Rad Laboratories GmbH, München, Germany). Samples containing 20 μg of protein were boiled for 5 min and loaded onto a 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis according to Laemmli (15). Then proteins were transferred to a polyvinylidene fluoride membrane by semi-dry blotting. The filters were incubated for 1 hr with blocking solution (triethanolamine-buffered saline supplement with 5% w/v dry milk powder) to block unspecific binding. Thereafter, filters were incubated for 1 hr with polyclonal rabbit-anti-HO-1 antibody (StressGen, Victoria, Canada) or polyclonal rabbit-anti-heat shock protein (HSP)-70 antibody (Santa Cruz, Heidelberg, Germany). After extensive washing with triethanolamine-buffered saline, 0.05% Tween 20, and 5% milk powder, filters were incubated with horseradish peroxidase-conjugated goat anti-rabbit immunoglobulin-G antibodies for 30 min. Antibody binding was visualized by chemiluminescence on BIO-MAX Kodak film.

Reverse-Transcriptase Polymerase Chain Reaction

Snap-frozen tissue samples from kidneys were homogenized using a Polytron homogenizer (IKA Labortechnik/Fischer Scientific).

Total RNA was isolated using Trizol reagent (GIBCO BRL, Grand Island, NY) according to the manufacturer’s instructions. Two micrograms of total RNA were reverse transcribed into cDNA by oligo-dT and random hexamer priming (Perkin Elmer, Weiterstadt, Germany). cDNA was used as the substrate for semiquantitative polymerase chain reaction (PCR). Primer sets were designed on the basis of published cDNA sequences or were those that have been used in published studies. The sequence of the primers, expected PCR product lengths, and annealing temperatures are as follows: tumor necrosis factor (TNF)-α forward GGT GAT CGG TCC CAA CAA GGA, reverse CAC GCT GGC TCA GCC ACT C, annealing temperature 61°C, 173 base pairs (16); monocyte chemoattractant peptide (MCP)-1 forward ATG CAG GTC TCT GTC ACG, reverse CTA GTT CTC TGT CAT ACT, annealing temperature 55°C, 447 base pairs (17); and porphobilinogen deaminase (PBGD) forward CAA GGT TTT CAG CAT CGC TAC CA, reverse ATG TCC GGT AAC GGC GGC, annealing temperature 59°C, 135 base pairs (17). An equal volume of each cDNA solution was used for amplification in 25 μL of reaction mixture containing 2 μL cDNA; 1× PCR buffer (Perkin Elmer, Norwalk, CT); 1 μM of each dNTP (dATP, dCTP, dGTP, and dTTP; Perkin Elmer, Norwalk); and 1 μM of specific 5′ and 3′ primers, and 0.75 units of AmpliTaq DNA Polymerase (Perkin Elmer, Norwalk). PCR was performed with a T-Gradient Thermo cycler (Biometra, Hannover, Germany).

Amplification was commenced with incubation for 4 min at 95°C, followed by amplification cycles as follows: 1 min at 95°C, annealing temperature for 45 sec, 1 min at 72°C, and in the end 4 min at 72°C. For the housekeeping gene PBGD, amplification was commenced with incubation for 6 min at 95°C, followed by amplification cycles as follows: 20 sec at 95°C, 30 sec at 59°C, 7 sec at 73°C, and in the end 7 min at 73°C.

PCR products (10 μL) were subjected to a gel electrophoresis (4% agarose gel, Sigma, Munich, Germany), which contained 4 μg/μL ethidium bromide. The DNA bands were visualized under ultraviolet light using a Bio-Imaging analyzer (Bio Doc II/NT, Biometra), and densities of target DNA bands were measured by scanning densitometry with a Bio-Imaging analyzer (Bio Doc II/NT, Biometra). The ratio of the densities allowed absolute amounts of RNA from unknown samples to be calculated and expressed as a ratio to PBGD internal control. Negative controls were performed routinely by running PCR without cDNA to exclude false-positive amplification products. RNA extraction and reverse transcription were performed twice for each sample. To determine relative gene expression values, each PCR reaction was repeated at least twice from all obtained cDNAs. To exclude possible amplification of genome DNA, PCR was performed on RNA samples directly. No amplification products were observed under these conditions.

Statistical Analysis

All data are given as means ± standard error of mean (SEM) because multiple groups were compared. For comparison of means for different groups, the Mann-Whitney test with option for multiple comparisons was applied (StatsDirect 2.2.2, Aswell, UK). A P value of less than 0.05 was considered significant.



Brain-dead and ventilated, non–brain-dead animals did not differ in their MAP before the intervention (brain-dead rats, 124 mm Hg; ventilated rats, 120 mm Hg; P =not significant [NS]). Within 3 min after induction of brain death, MAP increased to 148% of base line, followed by a sudden decrease to 52%, and slowly increasing thereafter. There was no change of MAP in ventilated, non–brain-dead animals. After 4 hr, there was no difference in MAP in both groups (Fig. 1A). The decrease of MAP in brain-dead animals was accompanied by a significant decrease of RBF (Fig. 2A), which was not observed in ventilated, non–brain-dead animals.

Figure 1
Figure 1:
(A) Effect of brain death on blood pressure. Course of mean arterial pressure (MAP) during the experiment. (B) Effect of different dosages of dopamine on the course of MAP. MAP is expressed as percent of baseline (MAP before intervention). (Each group consisted of ≥5 animals; bd, brain dead; bd + dopamine 2, brain dead + dopamine 2 μg/kg/min; bd + dopamine 6, brain dead + dopamine 6 μg/kg/min; bd + dopamine 10, brain dead + dopamine 10 μg/kg/min; bd + dopamine 14, brain dead + dopamine 14 μg/kg/min; ventilated + dopamine 10, ventilated animals + dopamine 10 μg/kg/min.)
Figure 2
Figure 2:
(A) Effect of brain death on renal blood flow (RBF). Course of RBF during the experiment. (B) Effect of different dosages of dopamine on the course of RBF. RBF is expressed as percent of baseline (RBF before intervention). (Each group consisted of ≥5 animals: bd, brain dead; bd + dopamine 2, brain dead + dopamine 2 μg/kg/min; bd + dopamine 6, brain dead + dopamine 6 μg/kg/min; bd + dopamine 10, brain dead + dopamine 10 μg/kg/min; bd + dopamine 14, brain dead + dopamine 14 μg/kg/min; ventilated + dopamine 10, ventilated animals + dopamine 10 μg/kg/min.)

Different dosages of dopamine did not significantly affect MAP (P =NS) (Fig. 1B). However, with increasing dosages of dopamine, the fulminant increase and sudden decrease of RBF was abrogated in brain-dead animals (brain-dead animals vs. brain-dead + dopamine 10 or 14 μg/kg/min:P <0.05; ventilated vs. brain-dead + dopamine 10 or 14 μg/kg/min:P =NS). In contrast, RBF in ventilated, non–brain-dead animals was only marginally increased by dopamine (P =NS) (Fig. 2B).


The kidneys of brain-dead rats were characterized by a strong infiltration of monocytes and macrophages compared with ventilated rats (P <0.001). This phenomenon was dose dependently inhibited by dopamine (dopamine 2 μg/kg/min:P =NS; dopamine 6 μg/kg/min:P =NS; dopamine 10 μg/kg/ min:P <0.05; dopamine 14 μg/kg/min:P <0.001), whereas dopamine in ventilated rats further promoted monocyte influx (dopamine 10 μg/kg/min:P <0.05 vs. ventilated rats; P <0.01 vs. brain-dead rats + dopamine 10 μg/kg/min) (Fig. 3A and C).

Figure 3
Figure 3:
(A) Monocyte/macrophage infiltration (ED1) in kidneys of ventilated and brain-dead animals with or without treatment of different dosages of dopamine. Data are expressed as cells per field of view ± standard error of mean (SEM) (400× enlargement; naïve, untreated control animals; ventilated, ventilated animals; bd, brain dead; bd + dop 2, brain dead + dopamine 2 μg/kg/min; bd + dop 6, brain dead + dopamine 6 μg/kg/min; bd + dop 10, brain dead + dopamine 10 μg/kg/min; bd + dop 14, brain dead + dopamine 14 μg/kg/min; ventilated + dop 10, ventilated animals + dopamine 10 μg/kg/min). Major histocompatibility complex (MHC) II expression in kidneys of ventilated and brain-dead animals with or without treatment of different dosages of dopamine. Data are expressed as cells per field of view ± SEM (400× enlargement; terms described in A). (B) Renal monocyte infiltration (ED1 staining). Sections of a 400× enlargement of groups are as follows: (a) naïve animals (unmodified controls), (b) brain-dead animals, (c) ventilated animals, (d) brain-dead animals + dopamine 2 μg/kg/min, (e) brain-dead animals + dopamine 10 μg/kg/min, and (f) ventilated animals + dopamine 10 μg/kg/min.

Similar, but not as striking, findings were observed with regard to renal MHC II expression determined as a marker of inflammation. Kidneys of brain-dead animals displayed a higher MHC II expression compared with kidneys of ventilated animals (P <0.05). At the dosage of 14 μg/kg/min dopamine, this strong expression of MHC II was significantly reduced (P <0.001; dopamine 10 μg/kg/min:P =NS; dopamine 6 μg/kg/min:P =NS; dopamine 2 μg/kg/min:P =NS). Dopamine treatment in ventilated rats did not alter MHC II expression (P =NS vs. ventilated rats) (Fig. 3B).

P-selectin, a cell-surface glycoprotein responsible for early recruitment of leukocytes, is rapidly expressed on vascular endothelium and platelets after inflammatory stimuli. Six hours after induction of brain death, there was a strong P-selectin expression in the renal vessels of brain-dead rats compared with ventilated rats (Table 1). Dose dependently, P-selectin expression was significantly down-regulated by dopamine, whereas in ventilated rats dopamine treatment did not alter P-selectin expression (Table 1).

Table 1
Table 1:
P-selectin expression in renal vessels

Western Blot

After 6 hr, there was a slight increase of HO-1 protein expression in the kidneys of brain-dead rats compared with ventilated rats. Because a dopamine dose of 10 μg/kg/min in brain-dead rats resulted in a significant reduction of monocyte influx with the least hemodynamic effect, we analyzed HO-1 protein expression in kidneys of brain-dead rats treated with this dose. Dopamine strongly up-regulated HO-1 protein in brain-dead rats, whereas HO-1 was only slightly increased in ventilated rats (Fig. 4). Dopamine treatment with 2 or 6 μg/kg/min did not significantly alter HO-1 expression in brain-dead rats (data not shown).

Figure 4
Figure 4:
Heme oxygenase (HO)-1 protein expression in ventilated and brain-dead animals with or without treatment of dopamine (10 μg/kg/min). (Western blot: 20 μg of protein of each probe was loaded onto a 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis gel.)

To further define whether HO-1 is an indicator for antioxidative properties or just a marker for cellular stress, we also analyzed the protein expression of HSP-70. In both groups (brain-dead animals and ventilated animals), dopamine did not significantly up-regulate HSP-70 expression (data not shown).

Reverse-Transcriptase Polymerase Chain Reaction

Renal TNF-α gene expression was significantly higher in brain-dead rats than in control animals (P <0.05). With dopamine treatment (10 μg/kg/min), this expression did not increase significantly in brain-dead rats (P =0.26), whereas dopamine promoted a significant increase of this inflammatory marker in ventilated rats (P <0.05) (Fig. 5A).

Figure 5
Figure 5:
(A) Tumor necrosis factor (TNF)-α gene expression in ventilated and in brain-dead animals treated with or without 10 μg/ kg/min of dopamine. (Reverse-transcriptase polymerase chain reaction [RT-PCR]: TNF-α gene expression is expressed as ratio to the “housekeeping” gene PBGD, mean ± SEM.) (B) Monocyte chemoattractant peptide (MCP)-1 gene expression in ventilated and in brain-dead animals treated with or without 10 μg/kg/min of dopamine. (RT-PCR: MCP-1 gene expression is expressed as ratio to the “housekeeping” gene PBGD, mean ± SEM.)

In comparison with kidneys of ventilated animals, kidneys of brain-dead rats displayed a significantly higher MCP-1 expression (P <0.05). This expression was not further up-regulated with dopamine treatment (10 μg/kg/min) (P =NS), whereas dopamine treatment in ventilated rats significantly increased renal MCP-1 expression (P <0.001) (Fig. 5B).


This study shows that dopamine in clinically relevant dosages down-regulates inflammation and enhances the antioxidative capacity of kidneys of brain-dead rats. Renal tissue of brain-dead animals was characterized by increased immunogenicity as delineated by strong macrophage infiltration, P-selectin, and MHC II expression. These characteristics were ameliorated by dopamine. In addition, dopamine prevented the up-regulation of TNF-α and MCP-1 gene expression, which are both associated with inflammatory response to tissue injury. Furthermore, our results indicate that dopamine increased HO-1, an enzyme that has been proven as antioxidative and cytoprotective in various settings of tissue pathology (18, 19).

In two independent retrospective clinical studies (5, 6), we clearly demonstrated that treatment of the brain-dead donor with dopamine is correlated with improved long-term kidney graft survival. So far, mostly negative effects of catecholamines on immediate graft function have been described in the literature (8, 9), but we reported beneficial effects on long-term outcome.

A strong monocyte invasion of kidneys of brain-dead rats, as seen in our model, has also been described by other groups (20). With an increasing dose of dopamine, this influx was significantly reduced. Similar findings were made with regard to renal MHC II expression. In contrast, dopamine further promoted monocyte influx in ventilated animals. This demonstrates a “high inflammatory state” of the kidney in brain-dead animals, which was ameliorated by dopamine. The high endothelial P-selectin expression in brain-dead animals also underlines an increased immunogenicity of the kidney. With the use of a similar model of brain death, Gasser et al. (21) reported that a recombinant soluble P-selectin ligand ameliorated renal injury induced during brain death. In our model, dopamine treatment significantly reduced P-selectin expression in renal vessels.

To further define the general inflammatory status of kidneys from brain-dead donors, we evaluated the renal gene expression of TNF-α. Although a significant up-regulation of TNF-α expression in brain-dead rats was not observed after dopamine treatment, renal TNF-α expression in ventilated rats significantly increased with dopamine treatment. Apparently the effects of dopamine on TNF-α mRNA are modified by brain death.

MCP-1 has been associated with progressive tubulointerstitial injury (22) and may therefore contribute to chronic allograft dysfunction. With a model of gradual onset of brain death in rats, Pratschke et al. (4) reported that MCP-1 was elevated in animals that had received a kidney transplant from brain-dead donors. In our model, kidneys of brain-dead rats were characterized by a high MCP-1 gene expression. Dopamine did not lead to a further significant up-regulation in this group, whereas it significantly increased MCP-1 expression in ventilated animals.

The role of HO-1 as an adaptive response and its possible immune modulatory function in tissues exposed to injurious stimuli has been well described (19, 23, 24). Recently, HO-1 has gained more attention for its cytoprotective properties during transplant rejection (18, 25, 26). Therefore, HO-1 may serve as a novel therapeutic target in organ transplantation. In our study, slightly increased HO-1 in brain-dead rats was strongly up-regulated with dopamine. It is unlikely that induction of HO-1 was mediated by dopamine receptors because it was previously demonstrated that the effect of dopamine on cultured endothelial cells was not influenced by the addition of a dopamine1- or dopamine2-receptor antagonist in an in vitro study by our group (13). Oxygen free radical scavengers, however, could block the HO-1 expression in vitro. Because expression of the “stress protein” HSP-70 was not significantly changed in both groups with the administration of dopamine, induction of HO-1 by dopamine will probably increase the antioxidative capacity in these kidneys.

It has been described that HO-1 is able to modulate endothelial cell adhesion molecules (27), and P-selectin expression in renal vessels was significantly reduced by dopamine in our study. As a possible consequence, there was a significantly lower monocyte infiltration in kidneys of brain-dead rats that had been treated with dopamine. This effect could also be directly mediated by HO-1, because Ishikawa et al. (28) described an inhibitory effect of HO-1 on monocyte transmigration.

In contrast with non–brain-dead animals, there was a high renal MCP-1 gene expression in brain-dead rats, underlining the high inflammatory status in these animals. But unlike in controls, dopamine treatment did not increase this chemokine in brain-dead animals. This effect could also be mediated by HO-1, because it has been reported that by HO-1 gene transfer MCP-1 levels were significantly depressed after hypoxic lung injury in mice (29). Conversely, Nath et al. (30) described high MCP-1 levels in HO-1 knockout mice challenged with oxidative stress. If the alterations in MCP-1 are mediated through HO-1, one would perhaps expect a significant decrease in MCP-1 in brain-dead animals after dopamine treatment; however, this was not observed. Whatever the mechanisms involved, dopamine treatment did not seem to have an unfavorable effect on MCP-1 gene expression.

In the two clinical studies cited, dopamine was mostly administered before the onset of brain death, reflecting clinical conditions. Because our hypothesis was that dopamine reduces the immunogenicity of kidney grafts of brain-dead donors, we started to administer dopamine with the onset of brain death in our animal model. To further define the best time point of dopamine administration (before or even after the onset of brain death), additional studies are warranted.

Apart from the inflammatory changes that we found in our brain death model, we also observed hemodynamic disturbances. These disturbances are known as “autonomic storm” in the literature. Although dopamine did not alter MAP in ventilated or brain-dead rats, it dose-dependently improved RBF in brain-dead animals. Although the mechanisms involved remain to be elucidated, the hemodynamic effects of dopamine on the kidney in brain-dead animals can also be regarded as favorable.


Dopamine exerts several “positive” effects on the kidneys of brain-dead animals. In contrast, dopamine further enhances markers of inflammation in ventilated, non–brain-dead animals. One possible explanation could be that in a “healthy” organism with intact systemic and local regulatory functions, the negative effects of dopamine predominate. In an organism that undergoes brain death with severe changes such as the “autonomic storm” and other mechanisms that are still incompletely understood, dopamine mobilizes the endogenous antioxidative capacity of renal tissue and mediates down-regulation of inflammation.

Our findings implicate that dopamine treatment in brain-dead organ donors may exert its positive effects on kidney grafts through several mechanisms. Further classification of the hierarchy of the pathophysiologic cascades involved may lead to a better understanding of the pathophysiology of brain death and ultimately to a more rational treatment of organ donors. A randomized, prospective clinical trial studying dopamine treatment of brain-dead kidney donors is in progress to supply the ultimate proof of the feasibility and efficacy of this approach.


We thank Susanne Meisinger for her excellent technical assistance.


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