The influence of artificially introduced N-glycosylation sites on the in vitro activity of Xenopus laevis erythropoietin.

Erythropoietin (EPO), the primary regulator of erythropoiesis, is a heavily glycosylated protein found in humans and several other mammals. Intriguingly, we have previously found that EPO in Xenopus laevis (xlEPO) has no N-glycosylation sites, and cross-reacts with the human EPO (huEPO) receptor despite low homology with huEPO. In this study, we introduced N-glycosylation sites into wild-type xlEPO at the positions homologous to those in huEPO, and tested whether the glycosylated mutein retained its biological activity. Seven xlEPO muteins, containing 1-3 additional N-linked carbohydrates at positions 24, 38, and/or 83, were expressed in COS-1 cells. The muteins exhibited lower secretion efficiency, higher hydrophilicity, and stronger acidic properties than the wild type. All muteins stimulated the proliferation of both cell lines, xlEPO receptor-expressing xlEPOR-FDC/P2 cells and huEPO receptor-expressing UT-7/EPO cells, in a dose-dependent manner. Thus, the muteins retained their in vitro biological activities. The maximum effect on xlEPOR-FDC/P2 proliferation was decreased by the addition of N-linked carbohydrates, but that on UT-7/EPO proliferation was not changed, indicating that the muteins act as partial agonists to the xlEPO receptor, and near-full agonists to the huEPO receptor. Hence, the EPO-EPOR binding site in X. laevis locates the distal region of artificially introduced three N-glycosylation sites, demonstrating that the vital conformation to exert biological activity is conserved between humans and X. laevis, despite the low similarity in primary structures of EPO and EPOR.

Introduction

Erythropoietin (EPO) is a hematopoietic cytokine that regulates the rate of red blood cell production [1, 2]. It binds to EPO receptors (EPORs) on the surface of erythroid progenitors, and then causes a preformed EPOR homodimer to undergo a conformational change which activates signal transduction pathways. The signal transduction events promote the survival, proliferation, and differentiation of erythroid progenitors, causing an increase in the number of circulating mature red blood cells. In humans and mice, EPO is a heavily glycosylated protein produced by adult kidneys, and is targeted to the bone marrow via blood circulation. The carbohydrate chain plays a part in the prevention of EPO being cleared from blood circulation, and therefore is vital for in vivo biological activity [3-10].

Erythropoiesis is one of the common functions widely conserved among vertebrates. Thus far, EPO sequences have been elucidated in at least 13 mammalian species [11-20], one amphibian (African clawed frog, Xenopus laevis) [21], and 3 teleost fish (pufferfish, Takifugu rubripes; zebrafish, Danio rerio; and goldfish, Carassius auratus L.) [22-25]. Interspecific comparison of the functional region can provide an insight into the universal structure-function relationships among EPO proteins. The functional region responsible for the receptor binding and biological activity of human EPO (huEPO) is described by several studies [26-30]. However, in other species, especially non-mammals, the functional region of EPO remains unexplored.

Previously, we described the cloning of X. laevis EPO (xlEPO) and its in vitro activity by a colony formation assay [21, 31]. Intriguingly, we found that xlEPO stimulates the proliferation of the cells that express human EPOR (huEPOR), even though the amino acid sequence of xlEPO is only 38% identical to that of huEPO [21]. This led us to expect that the functional region of EPO, which is responsible for in vitro activity, is conserved between X. laevis and humans. In light of this assumption, there are several findings that helped us to generate a working hypothesis. First, xlEPO has no N-glycosylation site [21], whereas huEPO has 3 N-glycosylation sites (at Asn24, Asn38, and Asn83) that are distal to the receptor-binding site [32]. Remarkably, these 3 N-glycosylation sites [13] and their biological cross-reactivity [33-36] are conserved in a number of mammalian EPOs. In addition, it has been observed that in huEPO glycosylation analogs with additional N-linked carbohydrates, the positions of the carbohydrate chains have an impact on whether the analog retains its in vitro biological activity [32, 37]. In light of these facts, we hypothesized that the activity of xlEPO should be retained, even if we introduce N-glycosylation sites to regions homologous to the 3 N-glycosylation sites of huEPO.

In this study, we tested this hypothesis by examining the activity of xlEPO muteins containing artificial N-glycosylation sites at positions homologous to those in huEPO (i.e., Asn24, Asn38, and Asn83). Those xlEPO muteins retained in vitro activity, demonstrating that the fundamental conformations of EPO-EPOR binding are conserved among humans and X. laevis; despite the low similarity in the primary sequences.

Notation of gene and protein symbols

The gene and protein symbols of Xenopus are suggested in Xenbase (Gene Nomenclature Guidelines; http://xenbase.org/gene/static/geneNomenclature.jsp). However, for reporting the comparative analysis of EPO in this study, we named the X. laevis erythropoietin protein “xlEPO” instead of xlepo.

Materials and Methods

Construction of N-glycosylated xlEPO muteins

N-glycosylated xlEPO muteins were constructed by introducing N-glycosylation consensus sequences (Asn-Xxx-Ser/Thr, where Xxx is any amino acid except Pro) into a region homologous to that of huEPO. Each N-glycosylation site results from the mutation of Thr24 Met26 to Asn24 Thr26, Asp38 Met40 to Asn38 Thr40, and Asp83 to Asn83 (Fig 1A and Table 1). N-glycosylated xlEPO cDNAs were produced by in vitro multiple site-directed mutagenesis of the xlEPO cDNA, as described below.

Fig 1

Design of N-glycosylated xlEPO muteins.

(A) Schematic drawings of Xenopus laevis erythropoietin (xlEPO) muteins. N- and O-glycosylation sites are indicated by closed and opened rhombuses, respectively. The terms to the left of the bars represent the names of the muteins. “X” in xlEPO-XXX indicates N-glycosylation positions (1, Asn24; 2, Asn38; 3, Asn83). (B) Amino acid sequences around the introduced N-glycosylation sites in the xlEPO muteins. The parallel sequences of wild-type xlEPO, human EPO, and mouse EPO are also shown. Consensus sequences for N-glycosylation (Asn-Xxx-Ser/Thr) are underlined, with the amino acid substitutions in bold.

Table 1

Primers used for multiple site-directed mutagenesis.

Primer	Oligonucleotide sequence (5′–3′)
Thr24Asn/Met26Thr	AGAGAATTGGAAAACGCCACGGACTCCTGCAAC a
Asp38Asn/Met40Thr	CAATTTCCTGAGAATATCACGGTCCCTGAAAC a
Asp83Asn	ATTTCATCTCCAATGCCAGCCTC a
tailed 5′-anchor	ACTTGCGGATCCATGGCGCTAGCCACCATGGGTGT b
tailed 3′-anchor	GCTAGTCTACCAACGTGA CGGTCGTTAACAGCTCT b
PCR 5′-anchor Fw	ACTTGCGGATCCATGGC
PCR 3′-anchor Re	AGAGCTGTTAACGACCG

The codons for mutated amino acids are underlined, with the point mutations in bold.

The start and stop codons are underlined and the unique nucleotide tail for subsequent mutant strand-specific PCR is in italics.

Fw, forward; Re, reverse.

Multiple site-directed mutagenesis and preparation of expression plasmids

Mutagenesis was performed as described by Seyfang and Jin [38], with some modifications. Double-stranded plasmid DNA containing the xlEPO gene [21] was used as a template. Mutagenic primers and tailed 5′- and 3′-anchor primers (Table 1) were phosphorylated before the annealing reaction. Two terminal-tailed primers with a unique 17-nucleotide tail (Table 1) were simultaneously annealed to the template DNA along with the set of mutagenic primers. Mutant strands were synthesized by primer extension and ligation in a single incubation step using T4 DNA polymerase and ligase. Full-length mutant strands were subsequently amplified by high-fidelity PCR with Pfu polymerase, which uses the unique mutant strand-specific tails introduced by the 2 terminal anchor primers. The specificity and band size of the amplified mutagenesis products were confirmed by 1.5% agarose gel electrophoresis. Following gel extraction and purification (QIAEX II Gel Extraction Kit; QIAGEN K.K., Tokyo, Japan) of the 584-bp band, the extracted DNA was cloned into a pGEM-T Easy vector (Promega K.K., Tokyo, Japan) and sequenced using an ABI3100 Genetic Analyzer (Applied Biosystems) and dye-terminator chemistry (BigDye Terminator; Applied Biosystems). Plasmids containing the desired mutations were digested with NheI and EcoRI. The DNA fragments were then cloned into the multiple cloning site (MCS) of the pIRES2-EGFP expression vector (BD Biosciences Clontech) containing the human cytomegalovirus immediate early promoter and an SV40 early polyadenylation signal sequence. pIRES2-EGFP contains the internal ribosome entry site (IRES) of the encephalomyocarditis virus between the MCS and the EGFP coding region; this enables both the xlEPO gene and the EGFP gene to be translated from a single bicistronic mRNA.

All constructs were transformed into E. coli DH5α. Several colonies were subsequently grown in Luria-Bertani broth containing 50 μg/mL of kanamycin. The plasmids were isolated using the alkaline-SDS lysis method.

Expression of xlEPO muteins in COS-1 cells

Monkey kidney fibroblast COS-1 cells (Cell Bank, RIKEN BioResource Center, Ibaraki, Japan) were transfected with vectors that incorporated the full coding sequence of the xlEPO genes. COS-1 cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% (v/v) FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin in a humidified atmosphere of 5% CO2 at 37°C. Cells (8 × 104 cells/well) were seeded in 24-well cell culture plates (Corning Incorporated, NY, USA). When the cultures reached 90% confluence, cells were transfected with a mixture of 0.8 μg plasmid cDNA and 1.5 μL Lipofectamine LTX (Life Technologies Corporation) per well in 100 μL Opti-MEM (Life Technologies Corporation) for 4 h at 37°C. After transfection, cells were incubated in antibiotic-free DMEM containing 10% FBS for 24 h. Cells were then incubated in serum-free, antibiotic-containing DMEM. After an additional 48 h, EGFP fluorescence was measured (excitation 485 nm, emission 528 nm) and conditioned media were collected. The collected media were arbitrarily concentrated using centrifugal filter devices (Amicon Ultra-4, 5000 NMWL; Millipore Corporation, Billerica, MA, USA) and stored at—80°C. Expression of the xlEPO muteins was confirmed by SDS-PAGE followed by Western blotting.

Gel electrophoresis and Western blot analyses

Samples were reduced by boiling for 5–10 min in sample buffer (50 mM Tris-HCl (pH 6.8), 20 mM dithiothreitol, 1% w/v SDS, 10% v/v glycerol, 2 mM EDTA, and 0.005% bromophenol blue) and subjected to SDS-PAGE with a separating gel containing 14% acrylamide. Protein markers (Biotinylated SDS-PAGE Standard Low Range; Bio-Rad Laboratories Inc.) were used for electrophoretic estimation of molecular weights. After electrophoresis, the proteins were transferred to a polyvinylidene difluoride membrane (Millipore). Protein transfer was carried out for 30 min with a constant current of 150 mA in a semi-dry electroblotter (Continental Lab Products Inc., San Diego, CA, USA) using anolyte (0.3 M Tris (pH 10.4) and 20% v/v methanol), transfer membrane (25 mM Tris (pH 10.4) and 20% v/v methanol), and catholyte solutions (25 mM Tris (pH 10.4) and 40 mM 6-amino-n-caproic acid, 20% v/v methanol). The blots were washed once with TBS (20 mM Tris-HCl (pH 7.5) and 500 mM NaCl) and then twice with TBS containing 0.1% (v/v) Tween 20 (TTBS). After blocking with 0.4% Block Ace (Snow Brand Milk Products Co., Ltd., Hokkaido, Japan) in TTBS for 45 min, each blot was incubated for 1 h with 2 μg/mL anti-peptide antibody against the xlEPO Thr44 to Ser57 sequence (anti-xlEPO sequence antibody) [31] or anti-GFP antibody (Medical & Biological Laboratories Co., Ltd., Nagoya, Japan) and then washed twice with TTBS. The blots were subsequently incubated for 1 h with biotinylated goat anti-rabbit IgG (1:6000; Bio-Rad), washed twice with TTBS, and treated with alkaline phosphatase-conjugated StrepTactin (1:6000; Bio-Rad) for 1 h. After 2 washes with TTBS followed by 2 more washes with TBS, proteins were detected by chemiluminescence (CDP-Star reagent; GE Healthcare Ltd., Tokyo, Japan). The bands were detected using a Luminescent Image analyzer LAS-3000 (Fuji Photo Film Co., Ltd., Tokyo, Japan). All washes were carried out for 5 min. All antibodies and alkaline phosphatase-conjugated StrepTactin were prepared in TTBS containing 0.4% Block Ace.

Physicochemical analysis of xlEPO muteins

Wild-type xlEPO and N-glycosylated muteins in mixed conditioned media were separated by reverse-phase high-pressure liquid chromatography (RP-HPLC) and cation exchange chromatography (CIEC). In RP-HPLC, the sample was applied at a flow rate of 0.4 mL/min to a YMC-Pack Protein-RP column (4.5 × 150 mm; YMC Co., Ltd., Tokyo, Japan) and separated with a linear gradient of 20%–60% acetonitrile in 0.1% trifluoroacetic acid over 30 min. A total of 10 fractions was collected once every 3 min (1.2 mL). In CIEC, the samples were adjusted to pH 2 with acetic acid, applied to 200 μL of SP Sepharose FF column (GE Healthcare), and eluted with 50 mM citrate/100 mM phosphate buffer (pH 2, 3, 4… 9). Each fraction was subjected to Western blot analysis.

De-N-glycosylation of xlEPO muteins

N-linked carbohydrates were removed by incubation with or without 0.8 U peptide N-glycosidase F (PNGase F; Roche Diagnostics K.K., Tokyo, Japan) in 20 mM sodium phosphate buffer (pH 8.6) containing 10 mM EDTA at 37°C for 24 h. Carbohydrate removal was assessed by SDS-PAGE and Western blot analysis.

Cell proliferation assay

Cell proliferation assays were performed as previously described [21]. Two cell lines were used: the interleukin-3-dependent murine cell line FDC/P2, which expresses the exogenous xlEPO receptor, xlEPOR-FDC/P2 [21], and the EPO-dependent human cell line, UT-7/EPO [39]. In the MTS assay, absorbance at 450 nm was measured using a microplate reader (POWERSCAN HT; DS Pharma Biomedical Co., Ltd., Osaka, Japan). Dose-response curves were fitted to a 4-parameter logistic model: y = (a—d) / {1 + (x / c) b} + d, where, a is E max and c is EC 50 [40]. Then, the E max (maximum effect) of the xlEPO mutein was determined without normalization by protein amount. The relative EC 50 (half-maximal effective concentration) of the xlEPO mutein was determined via normalization of v/v % of COS-1 supernatant EC 50 with its relative amount in COS-1 supernatant.

Statistical analysis

All experiments were performed at least 3 times with triplicate samples, and similar results were obtained each time. Error bars represent the standard deviations (SD). Either Student’s t-test or the Tukey–Kramer test was used, when appropriate.

Results

Design and expression of N-glycosylated xlEPO muteins

We introduced N-glycosylation consensus sequences into the xlEPO at positions homologous to those of huEPO (i.e., Asn24, Asn38, and Asn83) using in vitro mutagenesis (Fig 1A). The mutations were introduced both individually and in combinations. Thus, we constructed 7 different N-glycosylated xlEPO muteins with consensus sequences introduced at 3 different positions (Fig 1B). The wild-type xlEPO and N-glycosylated muteins were expressed in COS-1 cells using the lipofection method. All muteins were secreted from the COS-1 cells. The extent of glycosylation was evaluated by Western blot analysis using the anti-xlEPO sequence antibody (Fig 2A, top panel). The molecular weight of wild-type xlEPO was 18 kDa, which is equivalent to that of the peptide backbone. Western blots showed progressive increases in the molecular weights of xlEPO muteins from 18 to 22, 26, and 30 kDa, depending on the number of N-linked carbohydrates added. Treatment with PNGase F, which removes N-linked carbohydrates from glycoproteins, shifted the molecular weights of all muteins to 18 kDa, or the molecular weight of the wild type (Fig 2A, bottom panel). Thus, the increase in the size of the muteins in comparison to the wild-type xlEPO was attributed to N-linked carbohydrate content. Non-glycosylated Asn83 was observed in xlEPO-3, xlEPO-13, xlEPO-23, and xlEPO-123, which indicates that carbohydrate additions were efficient at Asn24 and Asn38, but inefficient at Asn83. The glycosylated xlEPO molecules showed 2 bands: a main band and a smaller one. These 2 bands were not observed after PNGase F treatment. This result shows that these 2 bands result from heterogeneity in the N-linked carbohydrate structure. The unsecreted protein was also examined by Western blot analysis (Fig 2B). Whole-cell lysates were loaded into each lane, and EGFP was used as a loading control to normalize the signals (Fig 2B, bottom panel). The wild-type xlEPO and N-glycosylated muteins left in the cell were detected (Fig 2B, top panel). To estimate the amount of xlEPO muteins in the COS-1 supernatant, we performed a densitometry analysis of Western blots. In Western blot analysis, the recognition of hyperglycosylated huEPO analogs by an anti-huEPO sequence antibody was reported to be weak compared with huEPO recognition, and was improved by the partial elimination of carbohydrate chains [41]. Therefore, we used Western blots of deglycosylated samples (Fig 2A, bottom panel) for densitometry analysis, because the added N-linked carbohydrates may prevent the anti-xlEPO sequence antibody from binding to the xlEPO muteins. The band intensity of wild-type xlEPO was defined as 1. The analysis revealed that the relative amount of xlEPO muteins in the COS-1 supernatant decreased depending on the number of N-linked carbohydrates added (Fig 2C). The detected amounts of the muteins, xlEPO-1, xlEPO-2, and xlEPO-3, containing 1 N-glycosylation site were approximately half that of the wild type. Moreover, the amounts of the muteins, xlEPO-12, xlEPO-23, and xlEPO-123, containing 2 or 3 N-glycosylation sites were only about one-quarter that of the wild type. Because the IRES-containing expression vector enables xlEPO and EGFP to be co-expressed from a single bicistronic mRNA, EGFP fluorescence intensity reflects xlEPO expression at the mRNA level. Therefore, the relative secretion efficiency of xlEPO muteins was obtained by dividing the relative amount (i.e. relative secretion level) by the relative expression level (Fig 2C). As a result, the secretion efficiencies of xlEPO muteins were revealed to decrease as the number of N-linked carbohydrates added increased, suggesting that glycosylation may have an adverse effect on xlEPO secretion.

Fig 2

Expression, secretion, and physicochemical properties of N-glycosylated xlEPO muteins.

(A) Western blot analysis of xlEPO muteins secreted from COS-1 cells. Conditioned media were concentrated 10-fold (top panel), and subjected to PNGase F treatment (bottom panel). Wild-type xlEPO and N-glycosylated muteins were separated by SDS-PAGE (reducing conditions) and visualized by Western blot analysis. Equal volumes of samples were loaded into each lane. (B) Western blot analysis of xlEPO muteins remaining in COS-1 cells. Whole-cell lysates were loaded into each lane. xlEPO (top panel) and EGFP (bottom panel) were visualized by Western blot analysis. Co-expressed nonsecretory EGFP was used as a loading control. (A, B) Standard proteins are shown at the far left, and the letters above each lane represent the names of the muteins shown in Fig 1. (C) Effects of N-glycosylation on the secretion of xlEPO muteins. The relative amounts of xlEPO muteins in the culture media of COS-1 cells were measured by densitometry analysis of Western blots (filled bars). The relative expression levels were calculated based on the fluorescence intensity of co-expressed EGFP (open bars). The relative efficiency of secretion was obtained by the division of relative amount by relative expression level (open circles). Values were normalized to that of the wild type, which was set at 1. *p < 0.05 and **p < 0.01 compared to the wild type. (D) Physicochemical properties of xlEPO muteins. Cation-exchange chromatography (CIEC, top panel) and the reverse-phase high-pressure liquid chromatography (RP-HPLC, bottom panel) of wild-type xlEPO and N-glycosylated muteins were performed as described in the Materials and methods section. The collected fractions were subjected to Western blot analysis. ‘N-CHO’ indicates the number of N-linked carbohydrates. Abbreviations are: EB, elution buffer; Fr., fraction.

Physicochemical properties of xlEPO muteins

The physicochemical properties of proteins, such as the electrical charge and hydrophilicity, affect their ligand–receptor binding affinity and/or protein stability in blood [42, 43]. To predict the isoelectric point (pI) of the xlEPO muteins, CIEC was performed (Fig 2D, top panel). Wild-type xlEPO was eluted at pH 8–9, and the xlEPO muteins containing 1 or 2 N-linked carbohydrates were eluted mainly at pH 7 or 8, respectively. Next, RP-HLPC was performed, in order to examine the hydrophilicity of the xlEPO muteins (Fig 2D, bottom panel). Wild-type xlEPO was eluted in fraction (Fr.) 9 at 52%–58% acetonitrile. Meanwhile, xlEPO muteins with 1 or 2 N-linked carbohydrates were eluted in earlier fractions (Fr. 8, 46%–52% acetonitrile or Fr. 7, 40%–46% acetonitrile, respectively). These results indicate that the xlEPO muteins are more acidic and hydrophilic than the wild type, owing to the negative charge of N-linked carbohydrates.

In vitro activity of xlEPO muteins

The effects of the added N-linked carbohydrates on the in vitro activity of xlEPO muteins were tested in cell proliferation assays using exogenous xlEPOR-expressing xlEPOR-FDC/P2 cells (Fig 3A), and huEPOR-expressing UT-7/EPO cells (Fig 3B). As expected, all xlEPO muteins stimulated the proliferation of both cell lines in a dose-dependent manner. Usually, dose-response curves as descriptors of ligand activity are characterized by differences in potency (described as EC 50) and efficacy (described as E max). Then, the relative potency and the relative efficacy of the xlEPO muteins on the proliferation of xlEPOR-FDC/P2 and UT-7/EPO cells were evaluated (Table 2 and Fig 4). As the number of N-linked carbohydrates increased, the relative potency of xlEPO muteins, which was obtained from the relative EC 50, tended to decrease in both cell lines (Fig 4A and 4B). On the other hand, the effect of N-linked carbohydrates on the relative efficacy of xlEPO muteins, which was obtained from E max, differed between the cell lines (Fig 4C and 4D). The efficacy on the proliferation of xlEPOR-FDC/P2 cells tended to decrease, as the number of N-linked carbohydrates increased (Fig 4C). More specifically, the efficacy of xlEPO-12 and xlEPO-23 was significantly lower than that of wild-type xlEPO, and xlEPO-1 and xlEPO-13 also showed a slightly lower efficacy compared to the wild type (Table 2). In contrast, the efficacy of the xlEPO muteins on the proliferation of UT-7/EPO cells was not significantly affected by the addition of N-linked carbohydrates (Fig 4D and Table 2). To confirm that the decrease in the activity of xlEPO muteins (EC 50 and E max) is attributed to the addition of N-linked carbohydrates, PNGase F-treated COS-1 supernatant was assayed by the proliferation of xlEPOR-FDC/P2 cells (Fig 3C). Removal of N-linked carbohydrates with PNGase F increased the in vitro activity of the xlEPO muteins (Fig 3C, middle and right panels), except xlEPO-3 (Fig 3C, left panel), when COS-1 supernatants were added at a final concentration of 6%. This indicates that the added N-linked carbohydrates decreased the in vitro activity of xlEPO.

Fig 3

Effects of wild-type xlEPO and the glycosylated muteins on the proliferation of X. laevis and human EPOR-expressing cells.

(A, B) The proliferative responses of xlEPOR-FDC/P2 cells (A) and UT-7/EPO cells (B) to the wild-type xlEPO and the N-glycosylated muteins were assessed using the MTS assay. Incubations were performed in triplicate, and the results are presented as the mean (± SD) from 1 of 4 independent experiments. (C) The proliferative responses of xlEPOR-FDC/P2 cells to wild-type xlEPO and the N-glycosylated muteins treated with (dashed line) and without (solid line) PNGase F were also assessed. The PNGase F-treated samples are the same fractions displayed in Fig 2A, bottom panel. Incubations were performed in triplicate, and the results are presented as the mean (± SD) from 1 of 3 independent experiments. **p < 0.01 for the comparison. Symbols are: multiple marks, wild-type xlEPO; open squares, xlEPO-1; open triangles, xlEPO-2; open circles, xlEPO-3; closed squares, xlEPO-12; closed triangles, xlEPO-13; closed circles, xlEPO-23; closed rhombuses, xlEPO-123; plus sign, Mock (control culture supernatant without xlEPO).

Table 2

Relative in vitro activities of xlEPO muteins.

	Relative EC 50 a (relative potency, %) b		Relative efficacy, % c
Protein	xlEPOR-FDC/P2	UT-7/EPO	xlEPOR-FDC/P2	UT-7/EPO
xlEPO-1	3.84 ± 0.54 (26)	1.34 ± 0.27 (74)	84 ± 15	106 ± 9
xlEPO-2	1.35 ± 0.79 (74)	5.17 ± 4.07 (19)	101 ± 5	108 ± 27
xlEPO-3	3.54 ± 1.50 (28)	2.37 ± 0.48 (42)	94 ± 6	100 ± 14
xlEPO-12	8.65 ± 1.58** (12)	3.24 ± 2.88 (31)	54 ± 16**	96 ± 8
xlEPO-13	4.47 ± 2.97 (22)	1.05 ± 0.43 (95)	89 ± 8	97 ± 20
xlEPO-23	3.25 ± 1.41 (31)	6.64 (15)	74 ± 14*	91
xlEPO-123	N.D.	187 (0.54)	N.D.	103

The relative half-maximal effective concentrations (EC 50) of the xlEPO muteins. EC 50 of wild-type xlEPO was defined as 1. Values are presented as mean ± SD of 2–4 independent experiments.

The relative potencies are expressed as the value of the wild type equal to 100%.

The relative efficacy of the xlEPO muteins. Values were obtained from maximum effects (E max) as the value of wild-type xlEPO equal to 100%. Values are presented as mean ± SD of 2–4 independent experiments.

*p < 0.05 and

**p < 0.01 for the muteins compared with the wild type.

N.D., not determined.

Fig 4

Relationship between carbohydrate content and biological activity.

The COS-1 supernatants containing the wild-type xlEPO and the N-glycosylated muteins were assayed by proliferation of xlEPOR-FDC/P2 cells (A, C) and UT-7/EPO cells (B, D). EC 50-derived relative potency (A, B) and E max-derived relative efficacy (C, D) are shown as a percentage relative to the wild-type xlEPO (Table 2) and are plotted against the number of added N-glycosylation sites. Each point represents different xlEPO muteins.

Discussion

In this study, we produced 7 different N-glycosylated xlEPO muteins and compared their biochemical properties of (i) secretion, (ii) physicochemical properties, and (iii) in vitro activity to those of the wild type.

The relative secretion efficiencies of the xlEPO muteins decreased depending on the number of N-linked carbohydrates added (Fig 2C). When the carbohydrate is added to the partially folded protein during translation [44], it has been hypothesized that the rates of synthesis or folding have a competitive relationship with the addition of the carbohydrate [45]. In light of this, the addition of carbohydrates was thought to reduce the secretion of xlEPO muteins at translation and/or during the secretion process.

Wild-type xlEPO was absorbed to the cation exchanger at pH 7 (Fig 2D). The pH of X. laevis plasma is approximately 7.4, the same as that of human plasma [46]. This indicates that wild-type xlEPO bears a positive electric charge in blood, whereas huEPO (pI, 4.4–5.1) bears a negative charge [47]. In case of huEPO, sialic acid of carbohydrates, which is negatively charged at physiological pH, is thought to decrease EPOR binding via charge repulsion. Accordingly, wild-type xlEPO is thought to easily access EPORs on cells compared to huEPO.

All N-glycosylated xlEPO muteins retained in vitro activity against the cell proliferation of both xlEPOR-FDC/P2 and UT-7/EPO (Fig 3A and 3B). The EC 50 and E max of a ligand represent the ability of the ligand to bind to its receptor (i.e., potency which reflects receptor-binding affinity) and its ability to cause an effect after binding to the receptor (i.e., efficacy which is intrinsic acticity), respectively. In the present study, we showed the relative EC 50 because we were able to roughly quantify the relative amounts of wild-type xlEPO and N-glycosylated muteins in conditioned media. As a result, we found that the addition of N-linked carbohydrates decreased the EC 50-derived relative potency of xlEPO muteins (Fig 4A and 4B). It has been reported that N-linked carbohydrates reduce the affinity of huEPO for its receptor, as well as the in vitro activity of huEPO [5]. This can be explained as follows: the negatively charged sialic acid and carbohydrate individually decrease the association rate constant of huEPO with huEPOR via their negative effect on intrinsic electrostatic enhancement [42]. It is also known that darbepoetin alfa, a hyperglycosylated huEPO analog with 2 extra N-linked carbohydrates, shows lower receptor-binding activity than huEPO [48]. Here, we confirmed that the N-glycosylated xlEPO muteins are more negatively charged than the wild type (Fig 2D). Taken together, it is thought that one of the reasons for reduced potencies of muteins is their lower receptor-binding affinities than that of wild type due to negative charge of carbohydrates.

Type of ligand effect on a receptor can be classified according to its behavior. Full agonists elicit the same level of full effect (i.e. E max, efficacy) as the natural ligand of the receptor, while partial agonists also cause an effect, but they cannot reach the same level of the natural ligand. The efficacy of the xlEPO muteins on xlEPOR-FDC/P2 cell proliferation was less than that of wild-type xlEPO (Fig 4C). In particularly, the efficacy of the xlEPO muteins containing N-linked carbohydrates on Asn24 was remarkably lower than that of wild-type xlEPO (Table 2). These data indicate that the muteins act as partial agonists against xlEPOR, and also suggest that the carbohydrate structure located at Asn24 interferes with xlEPOR binding. In contrast, the efficacy of xlEPO on the proliferation of UT-7/EPO cells was not changed by the addition of N-linked carbohydrates (Fig 4D). The E max of wild-type xlEPO and the N-glycosylated muteins was approximately the same as that of huEPO. These data indicate that the muteins act as near-full agonists against huEPOR, and also suggest that the N-linked carbohydrates at the Asn24, Asn38, and Asn83 sites of the xlEPO muteins do not interfere with huEPOR binding. Analysis of the effect of point mutations on bioactivity [26-29] and the crystal structure of huEPO, complexed with the extracellular ligand-binding domains of huEPOR [30], have identified amino acid residues important for receptor binding and biological activity. Twelve of the 18 amino acid residues that reduce the in vitro bioactivity of huEPO following point mutations, and interact with huEPOR residues, are conserved in xlEPO (6 of 10 in site 1, 6 of 8 in site 2), despite an only 38% overall sequence identity between xlEPO and huEPO (Table 3) [21]. These highly conserved amino acid residues are thought to allow xlEPO to cross-react with huEPOR and retain its proper orientation, avoiding interference from the added N-linked carbohydrates. With regard to xlEPOR, 7 of the 13 amino acid residues of huEPOR that interact with huEPO residues are conserved in xlEPOR (6 of 8 in site 1, 2 of 8 in site 2), which has an only 26% sequence identity with huEPOR in the extracellular domain (Table 3) [49]. Further investigation is needed to understand why the added carbohydrates on xlEPO interfere with the xlEPOR binding but not with the huEPOR binding.

Table 3

Amino acid residues of huEPO and huEPOR that contact each other in site 1 and site 2 intermolecular contact areas, and the corresponding residues in xlEPO.

Site 1				Site 2
xlEPO residue a	huEPO residue b , d	Neighboring huEPOR residues c , d	Conserved xlEPOR residues e	xlEPO residue a	huEPO residue b , d	Neighboring huEPOR residues c , d	Conserved xlEPOR residues e
K9	S9	H153		P5	L5	F93, S204	F85, S195
T10	R10	H153, E176		D8	D8	H153
D13	E13	P203		T10	R10	M150, S152, H153
I16	L16	P203, S204	S195	V11	V11	F93, M150, H153, S204	F85, S195
K17	L17	P203		M14	R14	L33, E34, P149, M150	L31
R20	K20	E202, P203		F15	Y15	S92, F93	F85
T44	T44	S92, F93, V94	F85	T78	Q78	A88
K45	K45	E62, S91, S92, F93, V94	F85	E91	D96	T87, A88
L46	V46	T90, S91, S92, F93	F85	K92	K97	E34
N47	N47	T87, A88, D89, T90, S91	D81	H94	V99	A88
V48	F48	L33, E34, S92, F93, F205	F85	S95	S100	T87, A88, T90, S91
G49	Y49	E34, T87		R98	R103	L59, E62, A88, D89, S91, V94	D81
N52	K52	E34		S99	S104	S91, S92, F93	F85
K121	R131	E60, D61, E62	E52, D53	H102	T107	F93, V94	F85
L123	I133	D61	D53	L103	L108	F93	F85
K130	K140	E60, D61	E52, D53	K105	R110	E60, D61	E52, D53
S133	R143	E60, P95	E52
N137	N147	F93, V94, P95, H114	F85
R140	R150	F93, H114, N116, E117, P203, S204	F85, N107, E108, S195
G141	G151	F93	F85
R144	K154	H153, S204	S195
L145	L155	F93	F85

xlEPO residues corresponding to right-hand “huEPO residues” [21]. Amino acid residues that are identical to huEPO residues are in bold.

Buried residues (>5.0 Å2) of huEPO in site 1 and site 2 intermolecular contact areas [30]. Available mutagenesis information [26–29] are highlighted according to the maximal degree of loss of in vitro bioactivity: bold underlined, >50-fold; bold, >5-fold; underlined, 2–5-fold; not highlighted, no effect.

HuEPOR residues within 4.5-Å distance from huEPO in the crystal structure of human EPO-(EPOR)2 complex (Protein Data Bank accession number, 1EER) [30]. Amino acid residues that are conserved in X. laevis are in bold [49].

Amino acid residues of huEPO and huEPOR that form interactions with each other, such as hydrogen bond and hydrophobic binding, are in italics [30].

Amino acid residues of xlEPOR that conserved in huEPOR[49].

huEPO, human erythropoietin; huEPOR, human EPO receptor; xlEPO, X. laevis EPO.

The most important finding presented here is that the carbohydrate structures, which are added to those xlEPO sites homologous to the N-glycosylation sites of huEPO, do not completely eliminate the in vitro biological activity of xlEPO, namely its binding to EPOR. This suggests that the functional region of xlEPO, which is responsible for its in vitro activity, must be located in a region other than the introduced N-glycosylation sites, as the 3 N-glycosylation sites of huEPO are distal to the receptor-binding site. We therefore conclude that a fundamental conformation of EPO-EPOR binding could be conserved between X. laevis and humans. Further point mutation study at the xlEPO-xlEPOR/huEPOR interface will give definite conclusions about their active sites. Furthemore, the key to understanding the observations presented here, as well as this conclusion, is the elucidation of the crystal structure of xlEPO in a complex with the extracellular domains of xlEPOR. Cross-species comparison of the tertiary structures of X. laevis and human EPOs (and other species such as fish) will facilitate the identification of the common denominators present in vertebrate EPOs.