Posttransplant erythrocytosis (PTE) was originally described in 1965 in a recipient of a living related renal allograft (1) who had undergone bilateral native nephrectomy prior to transplantation. Since then PTE has become a well-recognized complication of renal transplant occurring in 9–22% of allograft recipients (2–6). PTE is defined as an absolute hematocrit greater than 51%, a value that is at least 2.5 SDs higher than normal laboratory values for a healthy individual (2). This rise in hematocrit must be independent of other pathological conditions involving bone marrow or pulmonary disorders, presence of polycystic kidney or liver disease. The onset of PTE has been seen between 1–2 years after renal allograft transplantation, although it may happen at anytime (7–9). PTE can be transient and benign, however major thromboembolic events have been noted in 11.8–24% of PTE patients (10). The pathogenesis of PTE remains to be elucidated.
Some earlier studies raised the possibility that cyclosporin, an inhibitor of IL-2 that is implicated in inhibition of erythropoiesis, may be responsible for erythropoietin (Epo) independent erythremia (6, 11). However, if cyclosporin alone were indeed the culprit behind the pathogenesis of PTE, then its use would be expected to lead to erythrocytosis in other transplants (solid organ or bone marrow), which has not been reported.
More recent investigations have been targeted toward Epo, examining either the role of an abnormally elevated level of production (12–15), or increased hormone sensitivity of erythroid precursors (16, 17). The most likely source of elevated Epo would be the native kidneys and it was proposed that native nephrectomy would be a therapeutic maneuver in patients with PTE (12, 18). However, reports with respect to the level of Epo have been conflicting; 40–57% of patients with PTE have hormone levels undetectable by radioimmunoassay (7, 8, 19). This inconsistent finding of elevated serum Epo levels suggests that factors other than erythropoietin may be involved in the pathogenesis of PTE.
The attention has now shifted to the possible role of renin-angiotensin axis, specifically involving angiotensin II (AII) and its type 1 receptor (AT1R). Although anemia is not considered a side effect of angiotensin converting enzyme inhibitor (ACEI) use, some patients, typically renal transplant patients, being treated for hypertension with ACEI developed anemia (20, 21). This observation prompted evaluation of the beneficial efficacy of ACEI, which has now become an accepted modality of therapy for lowering hematocrit of patients with PTE. Although the Epo lowering ability of ACEI has been proposed as the mechanism of the hematocrit decrease (12–15), others have shown that the therapeutic effect of ACEI may involve inhibition of progenitor growth (16). The presence of the AT1R on BFU-E-derived cells has recently been reported (22). We determined that the expression of AT1R is increased on erythroid progenitors of patients with PTE when compared to renal transplant patients without PTE and normal volunteers. We also showed that this AT1R is functional. In contrast, expression of the EpoR on erythroid progenitors of renal transplant patients with and without PTE and normal volunteers was identical.
METHODS AND MATERIALS
The database of all renal allograft transplant recipients followed at the Milton S. Hershey Medical Center was reviewed. Patients with hematocrit of greater than or equal to 51% on two or more occasions were identified. Patients were chosen who were newly diagnosed, or were being treated with intermittent phlebotomy. Individuals with a history of polycystic kidney, hepatic dysfunction, pulmonary disease, bone marrow disorder, current use of ACEI or other cause for erythrocytosis including volume depletion were excluded. Renal transplant patients with normal hematocrit were randomly chosen as non-PTE controls, and were closely matched with respect to sex, age, weight, diuretic use, renal function, and immunosuppressive regimen. Normal healthy volunteers comprised the third group. Exclusion criteria were similar in all groups.
Preparation of BFU-derived erythroblasts.
Peripheral blood was obtained from all participants at the Milton S. Hershey Medical Center under protocols approved by the Institution’s Clinical Investigation Committee. A total of 60 ml of blood was obtained via routine venipuncture into heparinized syringes. Peripheral blood mononuclear cells were separated on Ficoll-Paque (Pharmacia, LKB Biotechnology, Inc., Piscataway, NJ) and cultured at 2×105 cells/ml in 0.9% methylcellulose media containing 30% fetal calf serum, 9.0 mg/ml deionized bovine serum albumin (Cohn fraction V; Sigma Chemical Co., St. Louis, MO), 1.4×10−4 M β-mercaptoethanol, and 2 U/ml erythropoietin (recombinant Epo>100,000 U/Mg; R & D Systems, Inc., Minneapolis, MN). The cultures were incubated in humidified 4% CO2 at 37°C. Single BFU-E, when cultured in methylcellulose, proliferate and differentiate over a period of 14 days to form large colonies containing 1–5×104 mature erythroblasts. These cells can be removed from culture at different days to study a well-defined population of normal human cells at distinct stages of maturation (23–25). Day 7 BFU-E-derived colonies consist of small numbers of nonhemoglobinized or poorly hemoglobinized blasts with a large proliferative ability and include a large number of erythroid colony forming units (CFU-E) (24). Day 10 cells are predominantly proerythroblasts and basophillic normoblasts and have a decreased proliferative capacity. Day 10 BFU-E-derived colonies are easily identified by light microscopy since they are partially hemoglobinized. Day 14 cells are terminally differentiating polychromophillic or orthochromic normoblasts. Cytocentrifuge preparations of aliquots of BFU-E-derived colonies routinely identified >99% as erythroid precursors.
Erythroid colonies were counted, harvested, and pooled in 2×106 cell aliquots on day 10 for the purpose of AT1 and Epo receptor quantification for each individual patient. The average number of cells per BFU-E-derived colony was determined.
Whole cell lysates were prepared by suspending 2×106 BFU-E-derived cells in cell lysate buffer [50 mM Tris HCl, pH8.0, 150 mM NaCl, 0.05% NP 40, 100 mM NaF, 1 mM EDTA, 1 mM EGTA, 0.08 mM phenylmethyl sulfonyl fluoride (PMSF), 0.01 mg/ml leupeptin, and 0.01 mg/ml aprotinin]. The suspension was vortexed and centrifuged at 10,000 rpm for 10 min. The supernatant was saved for Western blotting. Equal concentrations of total protein were mixed with equal volumes of 2X sodium dodeycyl sulfate (SDS) sample buffer (10% glycerol, 0.7 M β-mercaptoethanol, 3% SDS, 62 mM Tris, pH 6.8). This mixture was boiled for 5 min and separated on 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gel. Proteins were electroblotted onto Hybond-ECL nitrocellulose membrane (Amersham Life Sciences, Bucks, UK) according to the recommended procedures of the manufacturer. After blocking in 5% dry milk in TBST buffer [20 mM Tris Hcl, pH 7.5, 500 mM NaCl, 0.05% Tween-20 (Bio-Rad, Hercules, CA)], the membranes were cut into two pieces at the 46-kDa molecular weight. The half of the membrane with higher molecular weights (>46 kDa) was incubated with anti-AT1R (sc-579, Santa Cruz Biotechnology, Santa Cruz, CA; diluted 1:500) and the remaining half was incubated with anti-IκBα antibody (sc-847, Santa Cruz Biotechnology; diluted 1:50) overnight at 4°C. Donkey antirabbit antibody (1:1500 dilution) was used as the secondary antibody and membranes were detected with the ECL-Western blotting system (Amersham). The membranes containing the higher molecular weight proteins were stripped and reprobed with anti-EpoR antibody (sc-695, Santa Cruz Biotechnology; diluted 1:1000). Donkey antirabbit antibody (1:1500 dilution) was once again used as the secondary antibody and membranes were detected with the ECL-Western blotting system (Amersham).
Measurement of [Cai] with digital video imaging.
The functional assessment of AT1R was performed by stimulation of day 10 cells with angiotensin and observing the change in intracellular free calcium [Cai]. BFU-E-derived cells were removed from culture on day 10 and labeled with anti-human β2 microglobulin (Boehringer Mannnheim Corp., Indianapolis, IN) (26). Cells were bound to anti-mouse Ig-coated glass coverslips by incubating at 4°C for 40–60 min. The cells were then incubated in phosphate-buffered saline (PBS) at 37°C for 20 min with 2 μM Fura-2 acetoxymethyl ester (Molecular Probes, Inc., Eugene, OR). Total time lapse from removal of cells from culture to completion of Fura-2 loading was 3–5 hr. Cell viability as judged by trypan blue exclusion was >98%. Fura-2 loaded cells in PBS were visualized with the digital video imaging system previously described (26–28). By quantitating the fluorescence intensity ratio, R, at 350/380, small changes in [Cai] can be detected independent of local variations in cell thickness or dye content. Baseline and peak fluorescence intensity ratio, R, were measured with computer based digital video imaging system (26–28). Cells were stimulated either with Iscoves’s modified Dulbeco’s media (IMDM), Epo (2 U/ml), or AII (1×10−6 M). The dose of Epo was the optimal concentration for in vitro BFU-E cultures. The dose of AII was selected based on a dose response curve (not shown) and consistent with the growth response shown by Mrug et al. (22). IMDM containing 1% fetal calf serum (FCS) was used as control for nonspecific stimulation of [Cai] by protein. Previously, no increase in [Cai] was observed in erythroid precursors in response to a control hormone, insulin, chosen because insulin stimulates cell growth as well as caused an increase in [Cai] in nonerythroid cells (26).
All laboratory tests were performed through Milton S. Hershey Medical Center clinical laboratories. Hematocrit, mean corpuscular volume (MCV), and reticulocyte counts were measured using a Sysmex SE 9500 and Sysmex R 3000, respectively (Toa Medical Electronics co., Ltd., Kobe, Japan). Serum creatinine was measured using Roche Modular (Roche, Indianapolis, IN). Standard methods at indicated laboratories utilizing radioimmunoassay were used to determine serum erythropoietin (Geisinger Medical Center Laboratories, Geisinger, PA), angiotensin II (Nichols Institute Diagnostic, San Juan Capistrano, CA), and plasma renin activity (Hershey Medical Center, Hershey, PA). Specimen for laboratory analyses were collected simultaneously with blood used for culture.
Statistical analysis was performed using Prism (GraphPad Software Inc., San Diego, CA). The unpaired t test with Welch’s correction, where appropriate, was used to compare group means. Paired t test was used to analyze [Cai] data as well as comparison of pre- and posttransplant hematocrit. Nonparametric analysis was used to calculate Spearman r to assess any significant correlation amongst variables. All data are presented as mean±SEM, where appropriate. P <0.05 implies statistical significance.
Clinical and laboratory characteristics of posttransplant patients.
Table 1 shows the clinical characteristics of transplant patients with and without erythrocytosis. There were no statistically significant differences between the two groups with respect to age, sex, weight, immunosuppressive regimen, time on hemodialysis before transplant, or graft survival. Of all patients studied, only one NPTE patient and one normal donor were current smokers. There was an equal distribution of cadaveric versus living donor allografts. The use of diuretics was similar in the two groups. Two PTE and one NPTE patient had received combined kidney/pancreas allografts. Two other NPTE patients underwent retransplantation after a failed living donor allograft. None of the patients with PTE experienced graft failure. The graft survival time reported in Table 1 applies to the current functional graft. The mean time from transplant to onset of disease in this study was 41.8±15.4 months. This number is much higher than that reported in the literature; however, two of the patients developed PTE more than 10 years post transplant. Without those two patients, the mean was 21.6±8.7 months.
Table 2 shows the laboratory investigation of transplant patients with and without erythrocytosis. The hematocrit shown here and elsewhere in our report for PTE patients is the hematocrit at the time of study; in phlebotomized patients it was immediately before phlebotomy, not at diagnosis. In the normal (NORM) volunteers, only the hematocrit, MCV, and red blood cell (RBC) count were measured. There was a significant difference between PTE and NPTE patients with respect to hematocrit (51.0±0.9%, PTE; 40.7±0.9%, NPTE; 41.1±1.2%, NORM;P <0.0001) and number of RBC (5.7±0.2 M/μl, PTE; 4.6±0.1 M/μl, NPTE; 4.6±0.2 M/μl, NORM, P <0.0001). MCV was compared to rule out macrocytic or microcytic anemia and was found to be equivalent between groups. No differences were observed in serum creatinine, plasma renin activity, serum angiotensin II level, or blood pressure (Table 2). Although some studies have reported elevated serum erythropoietin levels in PTE patients, this was not observed in the current study.
In vitro growth of erythroid progenitors of posttransplant patients.
Culture data showed a statistically significant increase in the in vitro proliferation of BFU-E from patients with PTE compared to NPTE and normal controls (Figs. 1 and 2). Patients with PTE had a significantly increased number of BFU-E (48.9±2.3 per 2×105 mononuclear cells) versus NPTE (33.4±4.1, P =0.005) and NORM (35.4±5.0, P =0.02). Colony size was also significantly larger in cultures of patients with PTE (8280±1010 cells/colony) versus NPTE (4067±417, P =0.002) and NORM (3717±659.9, P =0.001). These results demonstrate the increased proliferative capacity of BFU-E from PTE patients and suggest that the increased erythroid proliferation seen in these patients is an intrinsic characteristic of their erythroid progenitors.
Expression of AT1R and EpoR on BFU-E-derived cells in postrenal transplant.
The expression of AT1R was first assessed in BFU-E derived cells at day 7, 10, and 14 (Fig. 3). A prominent band at 66 kDa was recognized by anti-AT1R antibody. This result was consistent with that reported in literature with this antibody (22) and was confirmed with western blotting with a second anti-AT1R antibody (AB 1525, Chemicon International Inc., Temecula, CA). A decrease in expression of AT1R was observed with the maturation of BFU-E-derived colonies. The antibody to IκBα, a ubiquitously present protein in day 7–14 cells, detected a band of 35–37 kDa. Antibody against the human erythropoietin receptor detected a band at approximately 66 kDa in BFU-E-derived cells but not in THP-1, a human monocytic leukemia line (ATCC TIB-202) used as control. Expression of EpoR and IκBα in differentiating BFU-E is also shown on Figure 3. Table 3 summarizes Western blot results of AT1R, EpoR, and IκBα expression in day 10 BFU-E-derived cells of patients with PTE, NPTE, and normal donors. Day 10 was chosen because erythroid colonies are easily identifiable due to their hemoglobinization and express measurable quantities of AT1R and EpoR. To control for variability in gel loading with whole cells lysates, densitometeric readings are reported as ratio of AT1R or EpoR over IκBα. AT1R expression, shown as a ratio over IκBα, was increased by 44% in patients with PTE (1.60±0.27) compared with NPTE renal transplant patients (1.11±0.17) (Table 3). A 32% increase in AT1R expression was also observed between PTE and NORM volunteers (1.21±0.17) (Table 3). These differences did not reach statistical significance, but sample size was small due to the number of available PTE patients untreated except for intermittent phlebotomy. However, a significant correlation was observed between the hematocrit and the level of AT1R expression (Fig. 4) for PTE patients (Spearman r =0.68, P =0.01). This correlation was not seen with the other two groups of subjects, NPTE (Spearman r =0.26, P =0.43), and NORM (Spearman r =0.43, P =0.22). In contrast, EpoR expression was essentially equivalent between all groups; PTE (0.60±0.11), NPTE (0.55±0.04), and NORM (0.58±0.07), P =NS.
Nonparametric regression analysis failed to reveal any significant correlation between BFU-E-derived colonies (number and size) and serum AII, Epo, or plasma renin activity. There were no significant correlations between the serum hormone levels mentioned above and the hematocrit for any groups. The relationship between hematocrit and serum Epo demonstrated no correlation for NPTE patients (Spearman r =0.11, P =0.74) and PTE patients (Spearman r =−0.4, P =0.2). Furthermore, there was no correlation observed between hematocrit and EpoR expression in PTE patients (Spearman r =0.52, P =0.07) or control groups; NPTE (Spearman r =0.07, P =0.82), Norm (Spearman r =0.39, P =0.26) (Fig. 4).
Response of AT1R on BFU-E to angiotensin II.
To assess the functional capability of the AT1R on erythroid precursors, individual day 10 BFU-E-derived cells were stimulated with angiotensin II. Stimulation of these single cells with erythropoietin served as a positive control and with IMDM as a negative control. There was a statistically significant increase in intracellular calcium in day 10 BFU-E-derived cells stimulated with either erythropoietin or angiotensin II (Fig. 5). The fluorescence intensity ratio (F350/F380=0.68±0.06) observed with Epo stimulation was 228% of baseline (F350/F380=0.30±0.01), expressed as 100% (P <0.0001, n=20). [Cai] after AII stimulation was 182% of baseline (0.29±0.01 to 0.52±0.04, P <0.0001, n=22), compared to a 116% change (0.25±0.2 to 0.29±0.03, P =NS, n=6) with IMDM. Thus, the AT1R is functional on BFU-E-derived cells.
This study, in which we report increased numbers of BFU-E and increased numbers of cells per BFU-E-derived colony in patients with PTE, confirms the findings of prior investigations (17, 22, 29). This observation has previously been attributed to increased sensitivity of erythroid progenitors to Epo (16). However, the Epo concentration used (2 U/ml) was higher than the Epo concentration at which Glicklich et al. (16) observed growth differences between PTE and control BFU-E. Thus increased sensitivity to Epo does not explain the results of our study. In contrast to previous reports of abnormal serum Epo level (12–15), no differences were observed here in Epo levels between patients with and without PTE. The plasma renin activity and serum AII levels were also similar between groups. Furthermore, no differences were noted here in EpoR expression in erythroid precursors of patients with and without PTE and normal volunteers. Therefore, the role of Epo as the sole causative agent in the pathogenesis of PTE is unlikely in the patients studied here.
Erythropoietin is a member of cytokine receptor superfamily, which share many signal transduction pathways that ultimately regulate gene expression, cell proliferation, and differentiation (30). It is a glycoprotein primarily produced by kidney, although the liver is an extrarenal site of production, in response to low oxygen saturation. Epo is obligatory for the proliferation and differentiation of erythroid cells. Epo interaction with its receptor rapidly induces dimerization/ oligomerization, which results in increased affinity for the Janus family of cytoplasmic tyrosine kinases, Jak2. Downstream signal transducers, which are subsequently activated, include STAT5, Ras, and IRS2 (31). The renin-angiotensin system primarily regulates blood pressure by maintaining fluid and electrolyte homeostasis. Recently, the role of angiotensin in erythropoiesis has become a subject of great interest. The activation of the AT1R by its substrate has been shown to activate Jak2/Stat, IRS2, and p70 S6 kinase pathways, common to activation of EpoR, in nonerythroid cells (32), raising the issue of whether AT1 and Epo receptor cross-talk on erythroid cells may have a functional role in erythropoiesis.
The mechanisms through which angiotensin may mediate an increase in erythropoiesis are poorly understood. Renin-angiotensin has been linked to erythropoiesis by previous investigators. Both renin and AII, when given to animals exogenously, have been shown to increase Epo production in the experimental animals (33–36). However, the effect of renin and angiotensin on the level of Epo in the hypoxic rat model (36), may be confused by the fact that hypoxia is also the major determinant of Epo synthesis and excretion. Julian et al. (37) showed an increase in serum Epo in humans with the administration of exogenous AII. These observations lead to the speculation that the therapeutic effect of ACEI lies in its ability to decrease Epo levels. However, many studies have now shown that ACEI lowers the hematocrit of PTE patients without affecting the serum Epo levels (10, 38, 39). An alternate theory of AII involvement may deal with Ca++ modulated cell growth. Our study showed the functional capacity of the AT1R on erythroid precursors by demonstrating an increase of [Cai] in single BFU-E-derived cells after AII stimulation, in a similar range as that observed with erythropoietin. A previous study showed (17) a 3-fold increase in the growth of BFU-E-derived cells when the Ca++ in the culture medium was increased although keeping the Epo concentration constant. Therefore, calcium may have a role in AII mediated erythropoiesis in an Epo-independent fashion. The role of Ca++ modulation in erythroid proliferation is currently under investigation (26–28).
Our data as well as other recent publications support involvement of the renin-angiotensin system in the pathogenesis of PTE. Angiotensin stimulation has previously been demonstrated to augment the growth of normal erythroid progenitors in culture (22). In addition, both ACEI and AT1R antagonists inhibit the growth of BFU-E- derived colonies (16, 22). Although these studies did not examine whether the effect of ACEI is caused by a direct interaction between the erythroid progenitor and ACEI, the ability of the AT1R antagonist losartan to reduce erythroid growth in vitro (22) and decrease the hematocrit of patients with PTE (37) suggests that the AT1R is involved. Here, pursuing the initial observation by Mrug et al. (22) that BFU-E-derived cells from normal donors express the AT1R at days 6 to 9 of culture, we examined the relative expression of AT1R on erythroid precursors. AT1R receptor levels were increased in BFU-E- derived cells of PTE patients but results did not reach statistical significance, probably due to small sample size. Nevertheless, the level of AT1R expression significantly correlated with the hematocrit in PTE, a finding that was not observed in control groups nor with EpoR. An increase in the sensitivity of AT1R for angiotensin II, which was not explored, might also be a potential pathway contributing to the pathogenesis of PTE (22, 37).
Thus, our evidence and that presented elsewhere supports the hypothesis that the AT1 receptor signal transduction pathway is involved in post transplant erythrocytosis and suggest a potential mechanism of action of ACEI. Both inhibitors of ACE and AT1 receptor antagonists, although through different mechanisms, may influence the ability of angiotensin II to interact with its receptor on erythroid precursors, down-modulating erythropoiesis. This effect may be more pronounced in the subgroup of patients with PTE, who have higher AT1R expression on erythroid precursors, resulting in a reduction in hematocrit. Although the pathogensis of PTE is likely to be multifactorial and involve more than an increase in AT1R, the AT1R signal transduction pathways activated by angiotensin II on erythroid cells need to be further investigated.
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