Advertisement
Browse Subject Areas
?

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Open Access

Peer-reviewed

Research Article

Isolation and Characterization of Renal Erythropoietin-Producing Cells from Genetically Produced Anemia Mice

  • Xiaoqing Pan,

    Affiliation Department of Medical Biochemistry, Tohoku University Graduate School of Medicine, Sendai, Miyagi, Japan

    ⨯
  • Norio Suzuki,

    Affiliation Department of Medical Biochemistry, Tohoku University Graduate School of Medicine, Sendai, Miyagi, Japan

    ⨯
  • Ikuo Hirano,

    Affiliation Department of Medical Biochemistry, Tohoku University Graduate School of Medicine, Sendai, Miyagi, Japan

    ⨯
  • Shun Yamazaki,

    Affiliation Department of Medical Biochemistry, Tohoku University Graduate School of Medicine, Sendai, Miyagi, Japan

    ⨯
  • Naoko Minegishi,

    Affiliation School of Nursing, Miyagi University, Sendai, Miyagi, Japan

    ⨯
  • Masayuki Yamamoto

    masiyamamoto@med.tohoku.ac.jp

    Affiliation Department of Medical Biochemistry, Tohoku University Graduate School of Medicine, Sendai, Miyagi, Japan

    ⨯

Abstract

Understanding the nature of renal erythropoietin-producing cells (REPs) remains a central challenge for elucidating the mechanisms involved in hypoxia and/or anemia-induced erythropoietin (Epo) production in adult mammals. Previous studies have shown that REPs are renal peritubular cells, but further details are lacking. Here, we describe an approach to isolate and characterize REPs. We bred mice bearing an Epo gene allele to which green fluorescent protein (GFP) reporter cDNA was knocked-in (EpoGFP) with mice bearing an Epo gene allele lacking the 3′ enhancer (EpoΔ3′E). Mice harboring the mutant EpoGFP/Δ3′E gene exhibited anemia (average Hematocrit 18% at 4 to 6 days after birth), and this perinatal anemia enabled us to identify and purify REPs based on GFP expression from the kidney. Light and confocal microscopy revealed that GFP immunostaining was confined to fibroblastic cells that reside in the peritubular interstitial space, confirming our previous observation in Epo-GFP transgenic reporter assays. Flow cytometry analyses revealed that the GFP fraction constitutes approximately 0.2% of the whole kidney cells and 63% of GFP-positive cells co-express CD73 (a marker for cortical fibroblasts and Epo-expressing cells in the kidney). Quantitative RT-PCR analyses confirmed that Epo expression was increased by approximately 100-fold in the purified population of REPs compared with that of the unsorted cells or CD73-positive fraction. Gene expression analyses showed enrichment of Hif2α and Hif3α mRNA in the purified population of REPs. The genetic approach described here provides a means to isolate a pure population of REPs, allowing the analysis of gene expression of a defined population of cells essential for Epo production in the kidney. This has provided evidence that positive regulation by HIF2α and negative regulation by HIF3α might be necessary for correct renal Epo induction. (282 words)

Citation: Pan X, Suzuki N, Hirano I, Yamazaki S, Minegishi N, Yamamoto M (2011) Isolation and Characterization of Renal Erythropoietin-Producing Cells from Genetically Produced Anemia Mice. PLoS ONE 6(10): e25839. https://doi.org/10.1371/journal.pone.0025839

Editor: Sean Bong Lee, The National Institute of Diabetes and Digestive and Kidney Diseases, United States of America

Received: June 17, 2011; Accepted: September 12, 2011; Published: October 11, 2011

Copyright: © 2011 Pan et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: This work was supported by Grants-in-Aid for Creative Scientific Research, Scientific Research on Priority Areas and Scientific Research from the Ministry of Education, Science, Sports and Culture Scientific Research from Japan Society for the Promotion of Science, Tohoku University, Global COE for Conquest of Signal Transduction Diseases with Network Medicine. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Erythropoietin (Epo) governs mammalian erythropoiesis. Epo is a glycoprotein hormone mainly produced in the kidney and liver in response to changes in tissue oxygen tension. Epo regulates erythropoiesis by supporting the survival of erythroid progenitors and stimulating their differentiation and proliferation in bone marrow, hence increasing the oxygen-carrying capacity of blood [1]. Lack of Epo during mouse development leads to lethality at embryonic day 13.5 (E13.5) due to severe anemia [2] and over- or under-production of Epo results in polycythemia or anemia clinically [1]. Epo production is considered to be controlled primarily at the level of gene transcription and Epo gene expression is strictly regulated in a tissue/cell-specific and hypoxia/anemia-induced manner [3]–[7].

Several tissues have been reported to express the Epo gene; but the ability to produce substantial amounts of Epo during hypoxia/anemia is restricted to the fetal liver and adult kidney [4]–[8]. The kidney plays a major role in oxygen sensing and contributes ∼90% of plasma Epo in adult animals [9]. However, difficulties in identification and purification of the renal Epo-producing cells (REPs) have limited the understanding of the mechanism for controlling Epo production in kidney. REPs are frequently reported to be peritubular fibroblast-like cells in kidney [6], [10], [11]; and a hypoxia-dependent Epo-producing cell line derived from human renal cancer was also described recently to exhibit fibroblast-like phenotype [12]. However, further details remain to be elucidated [5], [7], [13].

Current knowledge of the molecular mechanisms of oxygen-sensing and renal Epo gene expression has been extrapolated mostly from in vitro studies in hepatoma cell lines [14]–[16]. These studies have suggested that hypoxia responsiveness of the Epo gene depends on an enhancer containing hypoxia-responsive elements (HREs) located in the 3′ flanking region of the gene (3′ enhancer), to which the hypoxia-inducible transcription factor (HIF) 1 binds. HIF1 is composed of two subunits, HIF1α and HIF1β. HIF1β is constitutively expressed, but HIF1α expression, almost absent in normoxia, is increased during hypoxia. Under normoxic conditions, HIF1α is hydroxylated at two proline residues by specific prolyl-4-hydroxylases (PHD1–3) that allow the E3 ubiquitin ligase von Hippel-Lindau (pVHL) to bind to HIF1α and mark it for proteasomal degradation. In addition, HIF1α is regulated by the aspargine hydroxylase factor inhibiting HIF1 (FIH1), which inhibits p300/CBP (CREB-Binding Protein) binding to HIF1α. The activities of PHD and FIH1 are basically dependent on cellular oxygen concentration and thus qualify as cellular oxygen sensors. Low oxygen tension causes inactivation of PHDs and FIH1, allows HIFα to accumulate, forms active transcription factor-complex HIF with HIF1β, recruits transcriptional cofactors, and initiates the transcription of hypoxia responsive genes including the Epo gene. Thus the PHD/pVHL/HIF system likes to be the oxygen-sensing pathway regulating Epo gene transcription [17].

However, recent clinical and in vivo studies have suggested a new layer of complexity to the mechanisms involved in the cellular response to hypoxia/anemia. Evidence from mouse models and hereditary erythrocytosis in humans has revealed that HIF2α rather than HIF1α plays a vital role in oxygen-regulated erythropoiesis and renal Epo production is probably regulated by PHD2/pVHL/HIF2α pathway [13], [18], [19]. There are three HIFα family members: HIF1α, HIF2α, and HIF3α, which share a number of similarities e.g. DNA-binding sequence, oxygen-dependent hydroxylation. Unlike ubiquitously expressed HIF1α, expression of HIF2α and HIF3α is limited to several tissues [20]. Both HIF1α and HIF2α activate transcription, while HIF3α negatively regulates HIF1α and HIF2α activity [20]–[22]. There is no literature on HIF3α's role in hematopoiesis thus far.

In order to clarify the whole picture of Epo gene regulation, we have generated a panel of mouse lines. First, we genetically deleted the 3′ enhancer (referred to as the EpoΔ3′E allele) and showed that this enhancer is necessary for hepatic Epo expression during the perinatal period {E17–postnatal day 13 (P13)} but dispensable for renal Epo expression after birth. Mice homozygous for the targeted allele (EpoΔ3′E/Δ3′E) are viable and fertile, but exhibit anemia during late-embryonic and newborn stages [23]. Then, using a 180-kb Epo transgene with a green fluorescent protein (GFP) reporter (Epo-GFP), we recapitulated tissue-specific, hypoxia-inducible GFP expression in kidney and liver tissue of mouse. Mutation studies on the transgene indicated that GATA factors are required for suppression of ectopic expression of the gene, but not essential for the Epo gene induction in REPs [6]. Also, we developed GFP knock-in mice (EpoGFP/wt) by homologous recombination in mouse embryonic stem cells (NS and MY, unpublished data). By examining these mouse lines, we identified GFP-labeled REPs as a population of peritubular interstitial cells in the kidney after birth.

Taken together, all these data in vivo strongly imply novel mechanism(s) and necessitate detailed studies on REPs to explore a specific oxygen-sensing pathway underlying the hypoxia-induced Epo production in the kidney.

Fluorescence activated cell sorting (FACS) of GFP expressing cells has been widely used for the isolation of hematopoietic stem cells in our laboratory [24], [25]. To make the link between the molecular and cellular mechanisms of hypoxia-induced Epo expression, in this paper, we have addressed controversial issues in REPs using cell-sorting techniques. Our Epo-GFP transgenic mice are a source of REPs isolation, but a pretreatment to induce anemia is not always successful for stable GFP expression in the kidney [6]. We therefore generated EpoGFP/Δ3′E mice, in which REPs were labeled with GFP as a result of neonatal anemia caused by genetic modifications. Taking advantage of the strong GFP expression in anemic newborns carrying the 3′ enhancer deletion, we have purified by FACS a cell population responsible for anemia/hypoxia induced Epo expression in the kidney.

Results

Generation of EpoGFP/Δ3′E Mice

We first tried to define REPs by GFP fluorescence, which truthfully reflects endogenous Epo expression. We have established EpoGFP/wt mice by homologous recombination. In our EpoGFP/wt mice, the Epo locus was targeted by replacing the 3′ part of exon 2 through exon 4 with GFP (Figure 1A, middle panel). The mice homozygous for EpoGFP/GFP, which lacked a functional Epo gene, died around E13.5 from anemia, corresponding with the previous report on a Epo gene knockout mouse [2]. The heterozygous animals (EpoGFP/wt) were healthy and fertile; and distinct GFP expression, mimicking the endogenous Epo expression pattern, was observed in the kidney and liver under hypoxia/anemia conditions (NS and MY, unpublished data).

thumbnail
Download:
Figure 1. Generation of EpoGFP/Δ3′E mouse.

A Diagram shows structures of the wild type (wt, upper), EpoGFP (middle) and EpoΔ3′E (lower), respectively. IRES, internal ribosomal entry site. B,C An EpoGFP/wt heterozygote and an EpoΔ3′E/Δ3′E homozygote were bred to obtain the compound EpoGFP/Δ3′E genotype. EpoGFP/Δ3′E mice exhibited transient anemia during the perinatal stage: B at P5 looked pale; C presented a decreased Hct value even lower than that of EpoΔ3′E/Δ3′E at P4–P6 stage. The EpoΔ3′E/wt littermates were used as a normal control. *p<0.05.

https://doi.org/10.1371/journal.pone.0025839.g001

For steady Epo induction, we set out to genetically produce an anemia model by taking advantage of our established EpoΔ3′E/Δ3′E mice (Figure 1A, lower panel). As we previously reported in EpoΔ3′E/Δ3′E mice, deletion of the 3′ enhancer provokes transient anemia at late embryonic and neonatal stages due to defect in hepatic Epo production and erythropoiesis. This anemic phenotype is recovered in 2 weeks after birth when major Epo production site switches from the liver to kidney [23]. Crossing an EpoGFP/wt heterozygote with a mouse homozygous for EpoΔ3′E/Δ3′E allele, we generated EpoGFP/Δ3′E compound offspring. The compound mice basically showed a similar phenotype to that of their EpoΔ3′E/Δ3′E parents: anemia persisting after birth and recovered in the juvenile stage. EpoGFP/Δ3′E newborns were severely pale compared with EpoΔ3′E/wt littermates (Figure 1B). At P4–6, the Hematocrit (Hct) of the EpoΔ3′E/Δ3′E and EpoGFP/Δ3′E newborns were 22.7±5.3 and 18.0±2.0 (%), respectively (Figure 1C). At the same stage, the Hct of the EpoΔ3′E/wt control was 32.0±4.1%; this value is comparable with that of wild type (data not shown). Increased Epo mRNA could be detected in the kidney of the EpoGFP/Δ3′E newborns by quantitative (q) RT-PCR (see below).

Cellular distribution of GFP expression in anemic neonatal kidneys

We then examined newborn (P4–6) kidneys to see GFP expression by immunostaining studies. In microscopic observation of the kidney section with anti-GFP immunostaining, fluorescence was minimal in control kidney samples from P4–6 EpoΔ3′E/wt littermates, but was prominent in the kidneys from the EpoGFP/Δ3′E mice (Figure 2A,B). GFP fluorescence was also directly detected in fixed EpoGFP/Δ3′E kidney slices by confocal microscopy (data not shown). GFP signals were confined to the cortex-medulla junction and focally distributed along the curve of the kidney at P4–6 (Figure 2B). GFP-expressing cells were stellate-shaped, nesting around proximal tubules in the deep cortical labyrinth and outer strip area of the kidney (Figure 2C,D). These findings are consistent with previous descriptions [5], [6], [26], [27] and indicate that the GFP expression reflects the endogenous expression of the Epo gene in kidneys of anemic EpoGFP/Δ3′E newborns.

thumbnail
Download:
Figure 2. GFP expression in the anemic kidney of EpoGFP/Δ3′E mouse.

Representative photos of P5 neonates: kidney slices A from a nonanemic EpoΔ3′E/wt littermate as a negative control; B–D from EpoGFP/Δ3′E newborns. Light and confocal microscopy revealing the GFP signal concentrated in the deep-cortex∼outer-medulla region (B), where the Epo-producing peritubular interstitial cells are localized (C,D). Anti-GFP signals: green (Alexa 488), and dark-brown (diaminobenzidine). Scale bars: 25 µm. The sketch at the bottom illustrates peritubular localization of the GFP-expressing cells in the kidney.

https://doi.org/10.1371/journal.pone.0025839.g002

Cell type of the GFP-expressing kidney cells in EpoGFP/Δ3′E newborns

Kidney interstitium contains mainly fibroblastic, dendritic cells [28], and vascular endothelial as well as tubular cells have also been identified as the site of Epo gene expression in the kidney [5]. We therefore carried out marker studies on GFP-expressing kidney cells of P4–6 anemic newborns by dual immunostaining under confocal microscopy and FACS detection (Table 1). Expression of platelet-derived growth factor receptor β (PDGFRβ) (Figure 3C) and Ecto-5′-nucleotidase/CD73 (Figure 4), but not CD31 (Figure 3A), major histocompatibility complex class II (MHCII) (Figure 3B), or E-cadherin (Movie S1), indicated that REPs are cortical fibroblast-like, but not tubular, endothelial, or dendritic cells. Our three-dimensional (3D) movie reveals that REPs are tightly packed around tubules, but not tubular cells themselves (Movie S1). We also examined the expression of alpha-smooth muscle actin (α-SMA), a marker for the subtype of renal interstitial fibroblasts [28]. Staining with α-SMA antibody was observed in the medullar and cortical cells near the capsule, but did not overlap with GFP-positive cells (Figure 3D). Taken together, most neonatal REPs at P4–6 stage, are CD73-, PDGFRβ-positive, α-SMA-negative peritubular fibroblasts (Table 1). Thus, neonatal REPs have the typical characteristics of adult cortical fibroblasts previously reported in healthy rat kidney [29].

thumbnail
Download:
Figure 3. Cellular marker characterization of the GFP+ kidney cells of EpoGFP/Δ3′E mouse.

Dual immunostaining combined with confocal microscopy indicating GFP-expressing cells are not endothelial (CD31, A), dendritic (MHCII, B), and tubular cells (E-cadherin, Movie S1). GFP-positive interstitial cells express fibroblast-like markers (PDGFRβ, C), but not myofibroblast marker (α-SMA, D). Kidney slices from P5 EpoGFP/Δ3′E mice were stained with GFP (green) and the indicated antibodies directly labeled with phycoerythrin (PE, red) or indirectly labeled with Alexa 594 (red) respectively. Merged images of the same kidney section are shown. DAPI: blue/nucleus; Scale bars: 25 µm.

https://doi.org/10.1371/journal.pone.0025839.g003

thumbnail
Download:
Figure 4. Co-localization of GFP and CD73 in the peritubular interstitial cells of the EpoGFP/Δ3′E kidneys.

A–C Representitive images of confocal microscopy of kidney sections from P5 EpoGFP/Δ3′E mice. B A high-power view, demonstrated that GFP and CD73 were mostly co-localized in peritubular fibroblast cells (merged in yellow). Global merged view of the kidney sections from P5 EpoGFP/Δ3′E neonates (C), and EpoΔ3′E/wt littermates (D). D Under normal conditions CD73 showed a wide expression spectrum in a variety of cellular types including a few interstitial fibroblasts in the kidney. C During anemia note that CD73+ cells appeared to be increased mainly in the peritubular fibroblast population. GFP: green; CD73: red; DAPI: blue/nucleus; merged: yellow. Scale bars: 20 µm.

https://doi.org/10.1371/journal.pone.0025839.g004

thumbnail
Download:
Table 1. Expression of cellular markers in REPs by multiple immunostaining methods.

https://doi.org/10.1371/journal.pone.0025839.t001

CD73 is considered to be a reliable marker for identification of renal cortical fibroblasts [27], and Epo-expressing cells [6], [11], [12], [28], [30]. In the kidney sections of P4–6 EpoGFP/Δ3′E newborns, roughly 60%, but not all of GFP-positive cells, expressed CD73. Therefore, GFP-positive cells were divided into CD73+ and CD73− populations (Figure 4A–C). Both CD73+ and CD73− GFP-expressing cortical interstitial cells were remarkably increased in EpoGFP/Δ3′E neonates (Figure 4C) compared with in EpoΔ3′E/wt littermates (Figure 4D). In EpoΔ3′E/wt non-anemic kidney, only a few GFP-expressing cells constituting both CD73+ and CD73− were observed and located in the juxtamedullary layer of the cortex (Figure 4D).

Unlike GFP signals, which were restricted to peritubular interstitium, CD73 stains were widely spread in the renal cortex. Besides fibroblasts, proximal tubule (brush boarder), glomeruli (mesangial cells) and the cells in the medullar rays were also observed to express CD73 in sections. The proportion of CD73 expressing cells and their staining intensity in non-fibroblast cells were similar in both anemic and non-anemic kidney sections (Figure 4C, D), i.e. among various types of CD73-expressing kidney cells, only cortical fibroblasts showed a hypoxia/anemia-responsive tendency.

FACS sorting of GFP+ cells and Epo mRNA expression in the isolated cells

We isolated GFP+ cells from the kidneys of P4–6 EpoGFP/Δ3′E neonates by flow cytometry. Kidney cells from EpoΔ3′E/wt littermates were used as negative controls (Figure 5B, upper-left panel). The GFP+ population was present grossly with a low to intermediate GFP intensity and constituted up to 0.2% of the total fresh kidney cells (Figure 5B, upper-right panel). This yielded several thousands viable GFP+ cells per EpoGFP/Δ3′E mouse at P4–6 newborn stage, with a purity of greater than 75%. We also assessed the association of GFP-expressing cells with CD73. As shown in Figure 5B, lower-right panel, of GFP+ cells from P4–6 anemic kidneys, 63% were CD73+ and 37% were CD73−. A confocal microscopic image of the sorted GFP+ cells is shown with anti-GFP immunostaining (Figure 5C).

thumbnail
Download:
Figure 5. Isolation of REPs from the P4–6 EpoGFP/Δ3′E anemic kidneys.

A Cell purification protocol for a rare population of REPs from kidney. B, upper panel Representative FACS scatter plot of kidney cells from pooled P4–6 EpoGFP/Δ3′E neonates demonstrating GFP+ cells above the gate (set using EpoΔ3′E/wt control cells). B, lower panel Assessment by FACS of percentage of GFP+ and CD73+ fractions in the kidney cells from P4–6 anemic EpoGFP/Δ3′E neonates. Pooled results were from three independent experiments. Note CD73 expression divides GFP+ cells into two parts: CD73+ and CD73−. C Representative confocal microscopy of the FACS-sorted GFP+ cells (REPs) with anti-GFP immunostaining. Scale bar: 5 µm. D Analysis of relative Epo mRNA levels by qRT-PCR (Hprt as a loading control) in FACS-purified GFP+ or the remaining GFP− kidney cells from P4–6 EpoGFP/Δ3′E mice; CD73+ vs. CD73− fractionated cells were also evaluated. The data shown are from four experiments, each performed in duplicate. nd: not detectable; *p<0.05.

https://doi.org/10.1371/journal.pone.0025839.g005

Subsequently, we evaluated the expression of Epo mRNA in each sorted fraction from P4–6 EpoGFP/Δ3′E kidneys, and in unsorted kidney cells of P4–6 EpoΔ3′E/wt and EpoGFP/Δ3′E newborns as well. The hypoxanthine-phosphoribosyl-transferase (Hprt) gene was used as a loading control, because expression of HPRT is less affected by hypoxia/anemia [31]. As shown in Figure 5D, qRT–PCR revealed high Epo mRNA expression exclusively in samples from EpoGFP/Δ3′E animals including unsorted kidney cells, CD73−, CD73+ and GFP+ subsets. In EpoGFP/Δ3′E mice, compared with the unsorted kidney, Epo mRNA levels were ∼100 fold enriched in the GFP+ fraction, but not in the CD73+ fraction. Epo mRNA expression was low but detectable in EpoΔ3′E/wt kidney cells at P4–6, but not in the GFP-negative fraction of EpoGFP/Δ3′E kidney cells (Figure 5D). We have reported that neuronal markers are expressed by Epo-producing cells in the adult kidney [6]. Consistently, transcripts for microtubule-associated protein 2 (MAP2) and neurofilament light polypeptide (NFL) were detected in the GFP+ fraction from P4–6 EpoGFP/Δ3′E kidneys (data not shown). These results demonstrate our system for isolation of REPs is reliable and efficient.

Gene expression profile of oxygen-sensing and HIFs of the isolated REPs

We examined the expression of molecules, known to be involved in oxygen tension-dependent regulation [19], by qRT-PCR analysis. Compared to GFP-negative kidney cells, no enrichment of mRNA expression of oxygen sensor genes, Phd1–3 and Fih1 genes was found in the REPs fraction (Figure 6A). Hif2α but not Hif1α (Figure 6B, upper panel) mRNA expression was up-regulated in the REPs. In this line, no enrichment of HIF1α target genes [4], Pgk1 and Phd2, Phd3 were found in the REPs. These are consisted with recent reports on the relationship of HIF2α to renal Epo production [12], [29], [32], [33].

thumbnail
Download:
Figure 6. Expression profile of cellular oxygen-sensing and hypoxia-inducible molecules in the isolated REPs.

qRT-PCR analysis of the genes related to hypoxia response using RNA extracted from the sorted GFP− and GFP+ renal cells from P4–6 EpoGFP/Δ3′E mice. Hprt as a loading control *p<0.05. Data are for three experiments performed in duplicate. A Four genes related to cellular oxygen-sensing; B three Hifα isoforms. Also note that HIF1α targets: Pgk1, Phd2 are not enriched in the GFP+ fraction (REPs). C Three transcript variants of Hif3α in the purified GFP+ fraction (REPs), using a method for exponential PCR amplification at a fixed threshold (see also Materials and Methods).

https://doi.org/10.1371/journal.pone.0025839.g006

Hif3α mRNA expression was also enriched in the REPs ((Figure 6B, upper right panel). In the three alternatively spliced variants of Hif3α mRNA: Ipas, Nepas and Hif3α, Nepas is expressed during infantile and newborn stages, while Ipas is seen in adults. In accordance with a reported method [34], [35], we examined the expression of these splice variants in REPs, and detected all three variant mRNAs in the fraction of REPs at P4–6 stage. Nepas mRNA expression was the highest, suggesting Nepas might be a dominant form among the three splicing variants of the Hif3α at P4–6 infantile stage (Figure 6C).

Discussion

By generating EpoGFP/Δ3′E mice, we isolated a specific type of renal cells namely REPs, which are responsible for Epo production after birth. REPs 1) are fibroblast-like interstitial cells residing in the tubulo-interstitial compartment of the kidney in anemic hosts (Hct 18%); 2) constitute up to 0.2% of whole kidney cells. About 63% of REPs also express CD73, a marker for cortical fibroblasts and Epo-expressing cells in kidney [6], [10], [28], [30]; 3) highly express Hif2α, Hif3α, but not Hif1α mRNAs; 4) are efficiently isolated from naive kidney tissues as GFP-expressing cells in our mutant.

Isolation system of REPs

Previously, difficulties in the isolation of REPs prevented better understanding of the mechanisms of Epo regulation in response to hypoxia. We developed an isolation system that phenotypically labeled REPs in the kidney, by way of a GFP knock-in (EpoGFP) combined with a 3′ enhancer (EpoΔ3′E). This enabled us to purify REPs, a rare cell population, from kidney by FACS-sorting.

Our GFP knock-in strategy facilitates the capacity to express GFP based on endogeneous Epo gene expression confined to a rare population, without worries about aberrant or ectopic transgene expression. As a result, we identified REPs as peritubular fibroblast-like interstitial cells concentrated at the cortico-medullary junction, corresponding to our previous finding from the Epo-GFP transgenic mouse studies [5], [6].

In the history of Epo research, in situ hybridization is a classic method to localize Epo mRNA; but extensive studies lead to confusion of REPs' whereabouts: peritubular interstitium, or a tubular site [5]. Transgenic mice, created by integrating a marker gene with regulatory sequences of the Epo gene, have provided a powerful tool for accurate localization of REPs in the kidney [6], [8], [11]. In mice bearing an Epo/SV40 T antigen transgene, REPs were successfully identified but attempts to isolate REPs in vitro failed [8], [11]. Importantly, later transgenic mouse studies suggested that a much wider (20-kb∼) flanking region of the Epo gene was needed for adequate levels of transgene expression in kidney [5], [6], [36].

To induce Epo gene expression, pre-treatment to induce hypoxia/anemia is usually required. These procedures, such as bleeding or phenylhydrazine injection, are not always successful in inducing stable anemia. For instance, we have tried isolation of REPs using our Epo-GFP transgenic mouse [6]. The GFP+ population of the kidney cells from the transgenic mouse were not distinct under FACS detection, despite that severe anemia was induced by phlebotomy. EpoGFP/Δ3′E mice provide a handy and important source: lacking the 3′ enhancer demonstrated impaired hepatic Epo expression and profound anemia (Hct value was about 18% in newborn stage P4–6) and allowed us to directly sort REPs by FACS, which are constantly and stably labeled with GFP fluorescence in newborn kidneys. Moreover, considering the loose connections of renal tissues and the decreased interstitial volume of the cortex, renal tissues from newborns promised to be a better source of this rare cell population than adult kidney [37]. What is important that newborn REPs are fully functional with regard to Epo secretion, soon after birth [9], [23]. Indeed, newborn REPs (P4–6) displayed adult REPs phenotypes: fibroblast-like (CD73+/α-SMA−) [28], [29] with neuronal marker expression (MAP2+/NFL+) [6].

Cellular characters of the GFP-labeled REPs

GFP coupled with marker molecule analyses revealed that REPs are localized in the deep renal cortex, form a network around tubules and adjacent capillaries with their processes, and express fibroblast markers CD73, PDGFRβ and soluble guanylyl cyclase (sGC) (data not shown) but not other cell markers CD31, MHCII or E-cadherin. Surface expression of these molecules on REPs was verified by both confocal microscopy and flow cytometry. Therefore, all of these data support a widely accepted notion that REPs are peritubular fibroblast-like interstitial cells [6], [10], and resolves earlier conflicts on the cell identity in the literature [5].

Fibroblasts are considered to be an easy cell type to cultivate. However our attempts to culture REPs were not successful. It is interesting to consider that REPs are a type of fibroblasts in a resting state, and when induced to proliferate (signified by α-SMA expression), they lose their ability to produce Epo [38], [39], such as the case for renal fibrosis and renal anemia. Concurring with our previous finding that adult REPs express neuronal markers [6], we confirmed Map2 and Nfl mRNA expression in FACS-sorted REPs from P4–6 newborns. These observations further support the notion that REPs are unique fibroblast-like cells.

Association with CD73 during anemia

Our histological and flow-cytometrical examinations revealed that REPs constitute grossly 0.2% of total kidney cells, with 63% co-expressing CD73 in the case of our EpoGFP/Δ3′E mice (Hct 18% at P4–6 newborn stage). In the non-anemic kidneys, only a few REPs, composed of both CD73+ and CD73−, can be observed in the juxtamedullary layer of the cortex. These cells probably represent a basal level of Epo production under normal conditions, which is required for daily production of red blood cells in normal individuals. In anemic kidneys both CD73+ and CD73− REPs are robustly increased in a pattern that spreads outward from the deep cortex toward the capsule and the inner medullar.

qRT-PCR revealed a 100-fold enrichment of Epo mRNA in the GFP+ fraction, but no enrichment in the CD73+ fraction, compared with the unsorted total kidney cells. CD73 has a wide expression profile in renal cortex, occupying roughly 3% of the total kidney cells at P4–6 newborn stage based on our FACS study. In addition to the interstitial cells, such as fibroblasts, T-, B-lymphocytes, many parenchymal cells e.g. glomorular mesangial, proximate tubular (brush board), collecting duct cells etc. also express CD73 [28]. Comparing the pattern of CD73 staining in the anemic kidney section with the non-anemic one, it seems that anemia increases the number of CD73+ cortical interstitial fibroblasts but not other types of CD73+ cells. Because cortical fibroblasts are rare population, among all of the CD73+ cells, the percentage of CD73 fraction did not change much in anemic kidneys compared with non-anemic kidneys. The heterogeneity of CD73 expression in REPs may reflect different functional or matured stages during anemia [39]]. Recently, it has been reported that loss-of-CD73 does not affect the expression of Epo [40]. Our data that only a part of REPs express CD73 in anemic kidneys seem to conform this finding that the loss of CD73 has not impact on renal erythropoietin induction under hypoxia.

Expression profile of oxygen sensor molecules and transcriptional determinants in REPs

Clinically, congenital defects of the oxygen-sensing pathway have been reported including VHL, PHD2 and HIF2A mutations that cause secondary erythrocytosis through the EPO gene over-expression [1], [7]. PHD2 inactivation is sufficient to induce near max. renal Epo production [41], [42]; and recent RNAi-based studies confirmed the major role of PHD2 in Epo regulation in vitro as well as in vivo [43]. Human and rat PHD2 mRNA are hypoxically induced by HIFs for negative feedback regulation [44], [45]. In the carbon monoxide exposed rat, PHD3 protein was detected and co-localized with HIF2α in cortical interstitial cells of the kidneys [46]. Phd2, 3 are the targets of HIF1α [4]. In this study, we examined four genes encoding oxygen-sensor molecules (PHD1–3 and FIH1) and did not observe any enrichment in REPs compared with other cells of the anemic kidneys in our gene expression profiling. This may be because all of these four genes are ubiquitously expressed in kidney, with respect to various cell types of the kidney.

As described above, Hif2α, rather than Hif1α shows highly REPs-specific expression patterns at the mRNA level. Interestingly, Hif2α mRNA levels are particularly high in tissues that are important for the systemic delivery of oxygen, for example the lung, heart, endothelium and the carotid body [47]–[49]. Quite recently, HIF2α protein expression has been shown in the peritubular fibroblasts that express Epo and CD73 in rat kidneys [30]. Preferential binding of HIF2α protein to the HRE within the native Epo gene 3′ enhancer has been also confirmed in hepatocytes [50]. As renal Epo expression does not depend on the Epo gene 3′ enhancer [23], the existence of a possible renal enhancer with a different HRE awaits investigation.

Enrichment of Hif3a mRNA was also observed in the REPs. Transcripts of all of the three splicing variants (Nepas, Ipas and Hif3a) of Hif3a could be detected in newborn REPs (P4–6), where Nepas seems to be the dominant form of the three. Nepas and Ipas have been demonstrated to be hypoxia-induced factors due to the presence of functional HREs upstream of Exon1a, and act as negative regulators of the HIF pathway. Both Ipas and Nepas show a cell-, and stage-specific expression pattern [21], [22]. IPAS (inhibitory PAS protein) has already been reported to work as a negative feedback factor in a hypoxic condition in the cornea [21], [51], but there is no literature on its roles in hematopoiesis so far. Our targeted Hif3a knockout mice (Hif3a−/−) show an impaired cardiovascular formation around birth. This phenotype is possibly caused by over-expressed Endothelin-1 in pulmonary endothelium. HIF3α (Nepas) was suggested to suppress HIF2α-driven transcription of Endothelin-1 according to the localization and reporter assays [22].

We were curious to see if a similar mechanism exists in Epo gene regulation. We are starting to explore the function of HIF3α in erythropoiesis by examination of our established Hif3a−/− mice [22]. Hif3a−/− mice were viable and fertile without abnormalities under normal conditions. Based upon our preliminary data in mouse hypoxia experiments, it appeared that Epo transcript showed up-regulated tendency in Hif3a−/− kidneys, in contrast to the wild type counterparts. In a recent review, McIntosh et al. also mentioned an erythropoietic phenotype in their independent Hif3a−/− mice [52]. We, therefore, hypothesized HIF3α-related negative regulation is also necessary in renal Epo production during hypoxia/anemia. By this, homeostasis of red blood cell mass might be maintained to prevent erythrocytosis and thrombosis occurring in animals and human beings. HIF response to hypoxia is complex. A recent report has demonstrated that human HIF3A gene expression is induced by hypoxia through activation of HIF1α but not HIF2α [53]. It raises the possibility that in REPs, Hif3a mRNA expression might be up-regulated by HIF2α, because REPs preferentially express Hif2a rather than Hif1a.

Recently a renal cell line producing Epo with a hypoxia-dependent manner has been successfully established from a patient suffering from renal cancer [12]. Here, we report for the first time on isolation or purification of REPs in vivo. Our mouse enables the purification of a rare cell population specific for renal Epo expression during anemia and a detailed examination of the hypoxia-dependent aspect of the cells. Finally, we report the novel finding that Hif2α and Hif3α (but not Hif1α) mRNA are preferentially expressed in REPs. Combined with recent evidence in vivo about the role of HIF2α in erythropoiesis, we propose a hypothesis: positive regulation by HIF2α and negative regulation by HIF3α may be necessary for correct renal Epo induction during hypoxia/anemia.

Materials and Methods

Generation of EpoGFP/Δ3′E mice

All mice used were from a C57BL/6 genetic background and were strictly kept in the specific-pathogen-free conditions. All experiments were conducted in accordance with the regulations of The Standards for Human Care and Use of Laboratory Animals of Tohoku University. The protocol was approved by the Committee on the Ethics of Animal Experiments of Tohoku University (Permit Number: 21-Idou-144 and 22-Idou-113).

EpoGFP/Δ3′E mice were generated by mating mice heterozygous for EpoGFP/wt with mice homozygous for deletion of the 3′enhancer (EpoΔ3′E/Δ3′E) [23]. Genotyping was performed by polymerase chain reaction (PCR) with the primer sets listed in Table 2. From this mating, half of the offspring would be EpoGFP/Δ3′E mice. These mice are genetically deficient in Epo gene 3′enhancer activity and had anemia within two weeks after birth.

thumbnail
Download:
Table 2. Oligo-nucleotide sequences of primers used in this study.

https://doi.org/10.1371/journal.pone.0025839.t002

Hematological analysis

Whole blood was collected from the carotid arteries, and hematopoietic indices were measured using an automatic blood cell analyzer (Nihon Koden).

Immunostaining

Kidneys were immersion-fixed in 4% paraformaldehyde (Nakarai Tesque) for 3 hours at 4°C and embedded in OCT compound (Sakura Finetechnical). Frozen sections 20 µm in thickness were incubated with primary antibody for 16 hours at 4°C, and detected by Alexa Fluor 488 (Molecular Probes) or Alexa Fluor 555 (Molecular Probes) conjugated anti-IgG as second antibodies. Color detection was performed using diaminobenzidine as a chromogen (brown color staining). Nuclei were stained with 4′-diamidino-2-phenylindole (DAPI). Fluorescent images were observed using the LSM510 confocal imaging system (Carl Zeiss).

All of the primary antibodies were diluted 1∶500 in blocking solution (Dako) as follows: Anti-GFP (MBL); Biotinylated Anti-E-Cadherin (R&D systems); anti-CD31 (BD Pharmingen); phycoerythrin (PE) anti-CD73 antibody (BD Pharmingen); Alexa Fluor 647 Anti-CD73 (Biolegend); anti-α-SMA (Abcam); Allophycocyanin (APC) anti-CD140b (PDGFRβ) (Biolegend); APC anti-MHCII (eBioscience).

FACS analysis and cell sorting

EpoGFP/Δ3′E anemic newborns were sacrificed at P4–6. The kidneys were collected in PBS from several litters and teased away from their surrounding tissues. Single cell suspension was prepared using dispase (1.25 mg/mL; Invitrogen) in PBS–15% FCS for 60 min at 37°C followed by washing in DMEM-Ham's F-12–10% FCS and passing through a nylon mesh to remove any clumps. This cell preparation was approximately 85% single viable cells. Whole kidney was analyzed and sorted on the flow cytometer (FACSAria, Becton Dickinson). The effectiveness of each FACS separation was assessed by immediately resorting an aliquot of GFP+ and GFP− cells (data not shown). Greater than 75% of the GFP+ population resorted to the same gate used in the initial sort.

qRT-PCR analysis

Total RNA was extracted from FACS-purified cells using Isogen reagent (Nippon Gene), according to the manufacturer's protocol. RNA was then concentrated using RNeasy MinElute columns (Qiagen) and first strand cDNA synthesis was performed using the SuperScript III First Strand Synthesis System for RT-PCR (Invitrogen).

Primers for amplifying 100–300 bp of each PCR product were used (Table 2). PCR reactions were SYBR Green programmed and carried out using qRT-PCR Mastermix (Takara). Each sample was analyzed in duplicate or triplicate. The data were normalized by subtracting the difference of the CT values between the target genes of interest (Tgene) and that of Hprt mRNA, thereby obtaining a ΔCT (Tgene CT−HPRT CT). Relative expression (fold induction) was calculated as 2−(SΔCT−CΔCT) where SΔCT−CΔCT is the difference between the sample ΔCT (GFP+ cells) and the control ΔCT (GFP− cells). Both target gene and Hprt reactions approached 100% efficiency as determined by standard curves. PCR products were analyzed by dissociation curve and on agarose gels to check that a single band was amplified.

The molar ratio was calculated as previously described [33], [34]: molar ratio = [La×(1+Ea)CTa]/[Lb×(1+Eb)CTb]. La and Lb indicate lengths of the amplicon for 1a and 1b transcripts, respectively. Ea and Eb indicate the amplification efficiency of a primer set for 1a and 1b transcripts, respectively. CTa and CTb indicate the numbers of threshold cycles for the 1a and 1b transcripts, respectively.

Statistics

Statistical analysis was performed between samples and controls using t-test (two tailed, unequal variance, p≤0.05 cut-off).

Supporting Information

3D-movie of REPs and their spatial coordination, were made by compiling images collected using a Zeiss LSM 510 confocal microscope. One z-slice of the stack is shown in Movie S1.

Supporting Information

Movie S1.

3D image of REPs by confocal laser-scanning microscopy. Kidney sections from P5 EpoGFP/Δ3′E newborn were co-stained with anti-E-cadherin to label the tubular cells. GFP: green; E-cadherin: white; DAPI: blue/nucleus. Scale bar: 20 µm.

https://doi.org/10.1371/journal.pone.0025839.s001

(MOV)

Acknowledgments

We thank T. Moriguchi, M. Morita and T. Souma for scientific discussions, and H. Seto and E. Naganuma for technical assistance. We also thank Biomedical Research Core of Tohoku University Graduate School of Medicine and Tohoku University Center for Laboratory Animal Research for technical support. We are grateful to Chugai Pharmaceutical Co., LTD for comments and suggestions.

Author Contributions

Conceived and designed the experiments: XP NS MY. Performed the experiments: XP IH SY. Analyzed the data: XP IH SY. Contributed reagents/materials/analysis tools: XP NS IH SY. Wrote the paper: XP NS NM MY.

References

  1. 1. Fried W (2009) Erythropoietin and erythropoiesis. Exp Hematol 37: 1007–1015.
  2. 2. Wu H, Liu X, Jaenisch R, Lodish HF (1995) Generation of committed erythroid BFU-E and CFU-E progenitors does not require erythropoietin or the erythropoietin receptor. Cell 83: 59–67.
  3. 3. Stockmann C, Fandrey J (2006) Hypoxia-induced erythropoietin production: a paradigm for oxygen-regulated gene expression. Clin Exp Pharmacol Physiol 33: 968–979.
  4. 4. Jelkmann W (2007) Erythropoietin after a century of research: younger than ever. Eur J Haematol 78: 183–205.
  5. 5. Suzuki N, Obara N, Yamamoto M (2007) Use of gene-manipulated mice in the study of erythropoietin gene expression. Methods Enzymol 435: 157–177.
  6. 6. Obara N, Suzuki N, Kim K, Nagasawa T, Imagawa S, et al. (2008) Repression via the GATA box is essential for tissue-specific erythropoietin gene expression. Blood 111: 5223–5232.
  7. 7. Haase VH (2010) Hypoxic regulation of erythropoiesis and iron metabolism. Am J Physiol Renal Physiol 299: F1–13.
  8. 8. Maxwell PH, Ferguson DJ, Nicholls LG, Iredale JP, Pugh CW, et al. (1997) Sites of erythropoietin production. Kidney Int 51: 393–401.
  9. 9. Koury MJ, Bondurant MC, Graber SE, Sawyer ST (1988) Erythropoietin messenger RNA levels in developing mice and transfer of 125I-erythropoietin by the placenta. J Clin Invest 82: 154–159.
  10. 10. Bachmann S, Le Hir M, Eckardt KU (1993) Co-localization of erythropoietin mRNA and ecto-5′-nucleotidase immunoreactivity in peritubular cells of rat renal cortex indicates that fibroblasts produce erythropoietin. J Histochem Cytochem 41: 335–341.
  11. 11. Maxwell PH, Osmond MK, Pugh CW, Heryet A, Nicholls LG, et al. (1993) Identification of the renal erythropoietin-producing cells using transgenic mice. Kidney Int 44: 1149–1162.
  12. 12. Frede S, Freitag P, Geuting L, Konietzny R, Fandrey J (2011) Oxygen-regulated expression of the erythropoietin gene in the human renal cell line REPC. Blood 117: 4905–4914.
  13. 13. Wenger RH, Hoogewijs D (2010) Regulated oxygen sensing by protein hydroxylation in renal erythropoietin-producing cells. Am J Physiol Renal Physiol 298: F1287–1296.
  14. 14. Semenza GL, Nejfelt MK, Chi SM, Antonarakis SE (1991) Hypoxia-inducible nuclear factors bind to an enhancer element located 3′ to the human erythropoietin gene. Proc Natl Acad Sci USA 88: 5680–5684.
  15. 15. Pugh CW, Tan CC, Jones RW, Ratcliffe PJ (1991) Functional analysis of an oxygen-regulated transcriptional enhancer lying 3′ to the mouse erythropoietin gene. Proc Natl Acad Sci USA 88: 10553–10557.
  16. 16. Semenza GL, Wang GL (1992) A nuclear factor induced by hypoxia via de novo protein synthesis binds to the human erythropoietin gene enhancer at a site required for transcriptional activation. Mol Cell Biol 12: 5447–5454.
  17. 17. Semenza GL (2004) Hydroxylation of HIF-1: oxygen sensing at the molecular level. Physiology 19: 176–182.
  18. 18. Patel SA, Simon MC (2008) Biology of hypoxia-inducible factor-2alpha in development and disease. Cell Death Differ 15: 628–634.
  19. 19. Semenza GL (2009) Involvement of oxygen-sensing pathways in physiologic and pathologic erythropoiesis. Blood 114: 2015–2019.
  20. 20. Lendahl U, Lee KL, Yang H, Poellinger L (2009) Generating specificity and diversity in the transcriptional response to hypoxia. Nat Rev Genet 10: 821–832.
  21. 21. Makino Y, Cao R, Svensson K, Bertilsson G, Asman M, et al. (2001) Inhibitory PAS domain protein is a negative regulator of hypoxia-inducible gene expression. Nature 414: 550–554.
  22. 22. Yamashita T, Ohneda O, Nagano M, Iemitsu M, Makino Y, et al. (2008) Abnormal heart development and lung remodeling in mice lacking the hypoxia-inducible factor-related basic helix-loop-helix PAS protein NEPAS. Mol Cell Biol 28: 1285–1297.
  23. 23. Suzuki N, Obara N, Pan X, Watanabe M, Jishage KI, et al. (2011) Specific contribution of the erythropoietin gene 3′ enhancer to hepatic erythropoiesis after late embryonic stages. Mol Cell Biol 31: 3896–3905.
  24. 24. Minegishi N, Suzuki N, Yokomizo T, Pan X, Fujimoto T, et al. (2003) Expression and domain-specific function of GATA-2 during differentiation of the hematopoietic precursor cells in midgestation mouse embryos. Blood 102: 896–905.
  25. 25. Suzuki N, Suwabe N, Ohneda O, Obara N, Imagawa S, et al. (2003) Identification and characterization of 2 types of erythroid progenitors that express GATA-1 at distinct levels. Blood 102: 3575–3583.
  26. 26. Koury ST, Bondurant MC, Semenza GL, Koury MJ (1993) The use of in situ hybridization to study erythropoietin gene expression in murine kidney and liver. Microsc Res Tech 25: 29–39.
  27. 27. Eckardt KU, Koury ST, Tan CC, Schuster SJ, Kaissling B, et al. (1993) Distribution of erythropoietin producing cells in rat kidneys during hypoxic hypoxia. Kidney Int 43: 815–823.
  28. 28. Kaissling B, Le Hir M (2008) The renal cortical interstitium: morphological and functional aspects. Histochem Cell Biol 130: 247–262.
  29. 29. Marxer-Meier A, Hegyi I, Loffing J, Kaissling B (1998) Postnatal maturation of renal cortical peritubular fibroblasts in the rat. Anat Embryol (Berl) 197: 143–153.
  30. 30. Paliege A, Rosenberger C, Bondke A, Sciesielski L, Shina A, et al. (2010) Hypoxia-inducible factor-2alpha-expressing interstitial fibroblasts are the only renal cells that express erythropoietin under hypoxia-inducible factor stabilization. Kidney Int 77: 312–318.
  31. 31. Vengellur A, LaPres JJ (2004) The role of hypoxia inducible factor 1alpha in cobalt chloride induced cell death in mouse embryonic fibroblasts. Toxicol Sci 82: 638–646.
  32. 32. Scortegagna M, Ding K, Zhang Q, Oktay Y, Bennett MJ, et al. (2005) HIF-2alpha regulates murine hematopoietic development in an erythropoietin-dependent manner. Blood 105: 3133–3140.
  33. 33. Gruber M, Hu CJ, Johnson RS, Brown EJ, Keith B, et al. (2007) Acute postnatal ablation of Hif-2alpha results in anemia. Proc Natl Acad Sci U S A 104: 2301–2306.
  34. 34. Tanabe O, McPhee D, Kobayashi S, Shen Y, Brandt W, et al. (2007) Embryonic and fetal beta-globin gene repression by the orphan nuclear receptors, TR2 and TR4. EMBO J 26: 2295–2306.
  35. 35. Takayama M, Fujita R, Suzuki M, Okuyama R, Aiba S, et al. (2010) Genetic analysis of hierarchical regulation for Gata1 and NF-E2 p45 gene expression in megakaryopoiesis. Mol Cell Biol 30: 2668–2680.
  36. 36. Madan A, Lin C, Hatch SL, Curtin PT (1995) Regulated basal, inducible, and tissue-specific human erythropoietin gene expression in transgenic mice requires multiple cis DNA sequences. Blood 85: 2735–2741.
  37. 37. Sundelin B, Bohman SO (1990) Postnatal development of the interstitial tissue of the rat kidney. Anat Embryol (Berl) 182: 307–317.
  38. 38. Bechtel W, McGoohan S, Zeisberg EM, Muller GA, Kalbacher H, et al. (2010) Methylation determines fibroblast activation and fibrogenesis in the kidney. Nat Med 16: 544–550.
  39. 39. Muller GA, Strutz FM (1995) Renal fibroblast heterogeneity. Kidney Int Suppl 50S33–36.
  40. 40. Grenz A, Zhang H, Weingart J, von Wietersheim S, Eckle T, et al. (2007) Lack of effect of extracellular adenosine generation and signaling on renal erythropoietin secretion during hypoxia. Am J Physiol Renal Physiol 293: F1501–1511.
  41. 41. Kapitsinou PP, Liu Q, Unger TL, Rha J, Davidoff O, et al. (2010) Hepatic HIF-2 regulates erythropoietic responses to hypoxia in renal anemia. Blood 116: 3039–3048.
  42. 42. Minamishima YA, Kaelin WG Jr (2010) Reactivation of hepatic EPO synthesis in mice after PHD loss. Science 329: 407.
  43. 43. Fisher TS, Lira PD, Stock JL, Perregaux DG, Brissette WH, et al. (2009) Analysis of the role of the HIF hydroxylase family members in erythropoiesis. Biochem Biophys Res Commun 388: 683–688.
  44. 44. Epstein AC, Gleadle JM, McNeill LA, Hewitson KS, O'Rourke J, et al. (2001) C. elegans EGL-9 and mammalian homologs define a family of dioxygenases that regulate HIF by prolyl hydroxylation. Cell 107: 43–54.
  45. 45. D'Angelo G, Duplan E, Boyer N, Vigne P, Frelin C (2003) Hypoxia up-regulates prolyl hydroxylase activity: a feedback mechanism that limits HIF-1 responses during reoxygenation. J Biol Chem 278: 38183–38187.
  46. 46. Schodel J, Klanke B, Weidemann A, Buchholz B, Bernhardt W, et al. (2009) HIF-prolyl hydroxylases in the rat kidney: physiologic expression patterns and regulation in acute kidney injury. Am J Pathol 174: 1663–1674.
  47. 47. Tian H, McKnight SL, Russell DW (1997) Endothelial PAS domain protein 1 (EPAS1), a transcription factor selectively expressed in endothelial cells. Genes Dev 11: 72–82.
  48. 48. Wiesener MS, Jurgensen JS, Rosenberger C, Scholze CK, Horstrup JH, et al. (2003) Widespread hypoxia-inducible expression of HIF-2alpha in distinct cell populations of different organs. FASEB J 17: 271–273.
  49. 49. Tian H, Hammer RE, Matsumoto AM, Russell DW, McKnight SL (1998) The hypoxia-responsive transcription factor EPAS1 is essential for catecholamine homeostasis and protection against heart failure during embryonic development. Genes Dev 12: 3320–3324.
  50. 50. Rankin EB, Biju MP, Liu Q, Unger TL, Rha J, et al. (2007) Hypoxia-inducible factor-2 (HIF-2) regulates hepatic erythropoietin in vivo. J Clin Invest 117: 1068–1077.
  51. 51. Makino Y, Uenishi R, Okamoto K, Isoe T, Hosono O, et al. (2007) Transcriptional up-regulation of inhibitory PAS domain protein gene expression by hypoxia-inducible factor 1 (HIF-1): a negative feedback regulatory circuit in HIF-1-mediated signaling in hypoxic cells. J Biol Chem 282: 14073–14082.
  52. 52. McIntosh BE, Hogenesch JB, Bradfield CA (2010) Mammalian Per-Arnt-Sim proteins in environmental adaptation. Annu Rev Physiol 72: 625–645.
  53. 53. Pasanen A, Heikkilä M, Rautavuoma K, Hirsilä M, Kivirikko KI, et al. (2010) Hypoxia-inducible factor (HIF)-3alpha is subject to extensive alternative splicing in human tissues and cancer cells and is regulated by HIF-1 but not HIF-2. Int J Biochem Cell Biol 42: 1189–1200.