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. 2016 Feb;27(2):428-38.
doi: 10.1681/ASN.2014121184. Epub 2015 Jun 8.

Erythropoietin Synthesis in Renal Myofibroblasts Is Restored by Activation of Hypoxia Signaling

Tomokazu Souma  1 , Masahiro Nezu  1 , Daisuke Nakano  2 , Shun Yamazaki  3 , Ikuo Hirano  3 , Hiroki Sekine  3 , Takashi Dan  4 , Kotaro Takeda  5 , Guo-Hua Fong  5 , Akira Nishiyama  2 , Sadayoshi Ito  6 , Toshio Miyata  4 , Masayuki Yamamoto  7 , Norio Suzuki  8
Affiliations

Erythropoietin Synthesis in Renal Myofibroblasts Is Restored by Activation of Hypoxia Signaling

Tomokazu Souma et al. J Am Soc Nephrol. 2016 Feb.

Abstract

Erythropoietin (Epo) is produced by renal Epo-producing cells (REPs) in a hypoxia-inducible manner. The conversion of REPs into myofibroblasts and coincident loss of Epo-producing ability are the major cause of renal fibrosis and anemia. However, the hypoxic response of these transformed myofibroblasts remains unclear. Here, we used complementary in vivo transgenic and live imaging approaches to better understand the importance of hypoxia signaling in Epo production. Live imaging of REPs in transgenic mice expressing green fluorescent protein from a modified Epo-gene locus revealed that healthy REPs tightly associated with endothelium by wrapping processes around capillaries. However, this association was hampered in states of renal injury-induced inflammation previously shown to correlate with the transition to myofibroblast-transformed renal Epo-producing cells (MF-REPs). Furthermore, activation of hypoxia-inducible factors (HIFs) by genetic inactivation of HIF-prolyl hydroxylases (PHD1, PHD2, and PHD3) selectively in Epo-producing cells reactivated Epo production in MF-REPs. Loss of PHD2 in REPs restored Epo-gene expression in injured kidneys but caused polycythemia. Notably, combined deletions of PHD1 and PHD3 prevented loss of Epo expression without provoking polycythemia. Mice with PHD-deficient REPs also showed resistance to LPS-induced Epo repression in kidneys, suggesting that augmented HIF signaling counterbalances inflammatory stimuli in regulation of Epo production. Thus, augmentation of HIF signaling may be an attractive therapeutic strategy for treating renal anemia by reactivating Epo synthesis in MF-REPs.

Keywords: Pathophysiology of Renal Disease and Progression; anemia; chronic kidney disease; erythropoietin; fibrosis; hypoxia.

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Figures

Figure 1.
Figure 1.
Phenotypic changes of REPs upon injury. (A–F) Live imaging of REPs in the ISAM kidneys before (A–C) and 3 days after (D–F) ureteral obstruction (UO). Under a two-photon microscope, REPs and peritubular capillaries were detected with GFP expression (green) and rhodamine B-conjugated 70-kD dextran (red), respectively (B and C; high magnification views of A). Note that many processes from the cell body of REPs were attached along with the capillary walls at baseline (B and C), whereas some of the processes detached from the capillary wall (arrows in D) and attached to the tubular wall (arrowheads in E and F). Tubules (T) were visualized by the auto-fluorescence from the urine. (G) Quantification of the process directions of REPs. Numbers of directions either toward capillaries or toward tubules were counted. The data (mean±SD, n=3) indicate that the direction of the primary process of REPs changes toward the tubular side upon 3 days of UO. The number of process was not changed by the UUO procedure (normal REPs: 4.8±0.2 versus UUO-treated REPs: 5.3±0.4).
Figure 2.
Figure 2.
Kidney injury augments the hypoxic milieu. (A) Pimonidazole immunoblotting of whole kidney lysates. Pimonidazole-conjugated proteins were more abundantly accumulated in the injured kidneys by ureteral obstruction (UO) for 2 days than in the contralateral (Cont) or normal kidneys. α-Tubulin was used as a loading control. (B) Surface oxygen tension of UO-kidneys. Note that kidney surface oxygen tensions were reduced rapidly by UO. Green color indicates higher oxygen concentration than orange. (C) Pimonidazole immunoblotting of whole kidney lysates from ISAM (EpoGFP/GFP::TgEpo3′) and control littermates (EpoGFP/wt::TgEpo3′). Pimonidazole-conjugated proteins were accumulated in the kidney of ISAM due to the anemia, and the accumulation was enhanced by UO for 2 days compared with the contralateral kidneys (Cont). Labels #1 and #2 indicate different samples. α-Tubulin was used as a loading control. (D) Pimonidazole immunostaining of kidney sections. Hypoxic areas (brown, positive for pimonidazole adducts) of the ISAM kidney were widely spread throughout the cortical area after UO for 2 days (UO-kidney, right panels). Scale bar, 200 μm. (E) Mitochondrial membrane potential in injured kidneys of ISAM. TMRM, an indicator of mitochondrial membrane potential, was injected at each time indicated in the panels, and the fluorescent intensity (pseudo color) was measured by two-photon microscopy. The TMRM fluorescence in the tubular epithelial cells was high (orange to white) before obstruction (Pre), and the signals were decreased (dark blue to black) 4 hours after obstruction. Scale bar, 50 μm.
Figure 3.
Figure 3.
Hypoxic response is insufficient in injured kidneys. (A) HIF1α immunoblotting of whole kidney lysates from ISAM and the control littermates (EpoGFP/wt::TgEpo3′ genotype). HIF1α protein was accumulated in the kidney of ISAM due to the anemia, and the accumulation was reduced by ureteral obstruction for 2 days (UUO) compared with the contralateral kidneys (Cont). Labels #3 and #4 indicate different individual ISAM. β-Tubulin was used as a loading control. HIF2α was undetectable due to the technical difficulties. (B) Heat map diagram showing the changes of HIF-target gene expressions in the obstructed (UO-kidney) and normal kidneys of ISAM (from the microarray analyses; n=3). Red and green letters indicate genes upregulated and downregulated by UUO, respectively. (C) RT-qPCR analyses of the HIF target gene expressions in the obstructed (UO) and contralateral (Cont) kidneys of UUO-treated ISAM. The expression levels of the indicated genes (red asterisks in B) were measured 2, 7, and 14 days after UUO. Data from the sham-treated group were used as the starting point (0 day) and set as 1 (mean±SD). *P<0.05, **P<0.01 compared with the sham-treated group (n=3–5; one-way ANOVA with Tukey–Kramer test for multiple comparisons).
Figure 4.
Figure 4.
Deficiency of PHDs in Epo-producing cells leads to polycythemia. (A) Hct levels of mice harboring Epo-producing cell-specific deletion of the genes for PHD1, PHD2, PHD3, and/or HIF2α. Mice in which the Egln1 gene (encoding PHD2) was deleted in Epo-producing cells (P2-, P12-, P23-, P123-EKO) exhibited higher Hct levels than their control littermate mice (**P<0.01; n=4–17 for each genotype), and the polycythemic phenotype was abolished by deletion of the Epas1 gene (encoding HIF2α) in Epo-producing cells (P123H2-EKO). (B) Gross view of P123-EKO (Cre+) and the control (Cre–) littermate mice. Red cell mass (Hct) is shown in capillaries after centrifugation of the peripheral blood (left panel). P123-EKO mice exhibited red hind limbs (upper right panel) and larger spleens (lower right panel) compared with those of the control mice. (C–E) Spleen weight (C), plasma Epo levels (D), and Epo mRNA levels by RT-qPCR (E) were measured in the indicated genotype mice. Data from the contralateral kidneys of control mice were set as 100 in (E). *P<0.05, **P<0.01 compared with the Cre– controls (mean±SD, n=4–21 in each group; one-way ANOVA with Tukey–Kramer test for multiple comparisons).
Figure 5.
Figure 5.
PHD deficiency confers resistance to the loss of Epo-producing potential of REPs in injured kidneys. (A) RT-qPCR analyses of the Epo gene in kidneys of the indicated genotype mice at 2 days after UUO. Percentages of Epo mRNA levels in the obstructed kidneys compared with the contralateral kidneys (Cont) were calculated in each mouse after normalization with rRNA expression levels, and are shown as the mean±SD (n=4–8 in each group). *P<0.05, **P<0.01 compared with the control (Cre–) group (one-way ANOVA with Tukey–Kramer test for multiple comparisons). (B) High-level Epo mRNA expression in UO-kidneys of P123-EKO in later stages of fibrosis (UUO for 7 and 14 days). Epo mRNA levels were higher in the injured kidneys than in the contralateral healthy kidneys (Cont) in P123-EKO mice (Cre+). Data presentation is the same as described in (A). *P<0.05 compared with Cre– group (n=3–4 in each group). (C) Epo mRNA expressions of PHD-deficient kidneys (Cre+) underwent LPS challenge for 6 hours. Epo mRNA levels were analyzed by RT-qPCR. Declines of Epo mRNA level were compared with the vehicle-treated groups and shown as the mean±SD. *P<0.05 compared with the Cre– group (n=4, unpaired t test).
Figure 6.
Figure 6.
Activation of the Epo-gene transcription in PHD-deficient myofibroblasts. (A) Immunohistochemical detection of Epo-gene expressing cells in kidneys of EpoGFP/wt P123-EKO (Cre+; Egln2f/f::Egln1f/f::Egln3f/f::EpoGFP/wt::TgEpo-Cre genotype) and control (Cre–; Egln2f/f::Egln1f/f::Egln3f/f::EpoGFP/wt genotype) mice at 14 days after UUO. Frozen sections of the obstructed (UO) and contralateral (Cont) kidneys were stained with GFP (green: indicating transcriptional activity of the Epo-gene locus) and αSMA (red: marker for vascular smooth muscle cells [*] and myofibroblasts) antibodies, followed by 4',6-diamidino-2-phenylindole nuclear staining (blue). Original magnifications, x20. (B) Quantification of the number of Epo-GFP-positive cells in kidney sections. Data show mean±SD (n=3 in each genotype). Original magnifications, x100. (C) Higher magnification of A. GFP+ cells are present in the injured kidney of EpoGFP/wtP123-EKO mice and express αSMA at high (arrows) or low (arrowheads) levels. Scale bars, 100 μm in (A) and 20 μm in (B).
Figure 7.
Figure 7.
Renal inflammatory and fibrogenic response are not affected by PHD deficiency in REPs. (A) Picrosirius-red staining of the kidneys from P123-EKO (Cre+) and littermate control (Cre–) mice at 14 days after UUO. Note that collagen accumulation is stained in red. Scale bar, 100 μm. (B) Quantification of picrosirius-red positive area of (A). Data from the Cre– kidneys were set as 1 (mean±SD, n=3 in each group). (C) Expression profiles of fibrosis- and inflammation-related genes in the kidneys from P123-EKO (Cre+) and the littermate control (Cre–) mice upon UUO for 7 and 14 days. Data (mean±SD, 3 or 4 mice in each group) from RT-qPCR analyses were normalized with rRNA expression levels. *P<0.05 compared with the Cre– group (one-way ANOVA with Tukey–Kramer test for multiple comparisons).
Figure 8.
Figure 8.
Augmentation of HIF signaling restores the Epo-producing ability of renal myofibroblasts. (A) Schematic model of the Epo-gene regulation in REPs. Expression of the Epo gene is mainly regulated by the PHD2-HIF2α axis in REPs. Epo-gene transcription is inactive under normal oxygen (O2) conditions (OFF-REP). In REPs sensing hypoxia (ON-REP), oxygen-dependent PHD activity is blocked; thereby HIF2α is stabilized for activation of the Epo-gene transcription. Kidney injuries induce myofibroblastic transformation of REPs through the activation of inflammatory and fibrogenic signals and inactivate HIF2α through PHDs in spite of their worsened hypoxic milieus. Therefore, the inactivation of PHDs in MF-REPs leads to reactivation of Epo-gene expression. (B) Summary of the results from deletion of genes for PHD isoforms in REPs. This study proposes a therapeutic window (blue, around double deletion of PHD1 and PHD3) for inducing Epo production from diseased kidneys without polycythemic complications caused by PHD2 deletion.
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