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Blockade by phosphorothioate aptamers of advanced glycation end products-induced damage in cultured pericytes and endothelial cells
Microvascular Research 90 (2013) 64–70
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Brief Communication
Blockade by phosphorothioate aptamers of advanced glycation end products-induced damage in cultured pericytes and endothelial cells Yuichiro Higashimoto a,⁎, Takanori Matsui b, Yuri Nishino b, Junichi Taira a, Hiroyoshi Inoue c, Masayoshi Takeuchi d, Sho-ichi Yamagishi b,⁎⁎ a
Department of Chemistry, Kurume University School of Medicine, Kurume, Japan Department of Pathophysiology and Therapeutics of Diabetic Vascular Complications, Kurume University School of Medicine, Kurume, Japan Department of Chemistry, Keio University School of Medicine, Kanagawa, Japan d Department of Advanced Medicine Medical Research Institute, Kanazawa Medical University, Ishikawa, Japan b c
a r t i c l e
i n f o
Article history: Accepted 27 August 2013 Available online 5 September 2013
a b s t r a c t Advanced glycation end products (AGEs) not only inhibit DNA synthesis of retinal pericytes, but also elicit vascular hyperpermeability, pathological angiogenesis, and thrombogenic reactions by inducing vascular endothelial growth factor (VEGF) and plasminogen activator inhibitor-1 (PAI-1) through the interaction with the receptor for AGEs (RAGE), thereby being involved in the pathogenesis of diabetic retinopathy. In this study, we screened novel phosphorothioate-modified aptamers directed against AGEs (AGEs–thioaptamers) using a combinatorial chemistry in vitro, and examined whether these aptamers could inhibit the AGE-induced damage in both retinal pericytes and human umbilical vein endothelial cells (HUVECs). We identified 11 AGEs–thioaptamers; among them, clones #4, #7s and #9s aptamers had higher binding affinity to AGEs–human serum albumin (HSA) than the others. Surface plasmon resonance analysis revealed that KD values of #4s, #7s and #9s were 0.63, 0.36, and 0.57 nM, respectively. Furthermore, these 3 clones dose-dependently restored the decrease in DNA synthesis in AGE-exposed pericytes. AGEs significantly increased RAGE, VEGF and PAI-1 mRNA levels in HUVEC, all of which were completely blocked by the treatment with 20 nM clone #4s aptamer. Quartz crystal microbalance analysis confirmed that #4s aptamer dose-dependently inhibited the binding of AGEs–HSA to RAGE. Our present study demonstrated that AGEs–thioaptamers could inhibit the harmful effects of AGEs in pericytes and HUVEC by suppressing the binding of AGEs to RAGE. Blockade by AGEs–thioaptamers of the AGEs–RAGE axis might be a novel therapeutic strategy for diabetic retinopathy. © 2013 Elsevier Inc. All rights reserved.
Introduction Diabetic retinopathy is one of the most miserable complications in diabetes and is a leading cause of acquired blindness among the people Abbreviations: AGEs, advanced glycation end products; EC, endothelial cells; VEGF, vascular endothelial growth factor; PAI-1, plasminogen activator inhibitor-1; RAGE, receptor for AGEs; SELEX, systematic evolution of ligands by exponential enrichment; AGEs–thioaptamers, phosphorothioate aptamers directed against AGEs; HUVECs, human umbilical vein endothelial cells; HSA, human serum albumin; BSA, bovine serum albumin; ssDNA, single-stranded DNA; PCR, polymerase chain reactions; dATP(αS), 2′-deoxyadenosine-5′-O-(1-thiotriphosphate); dTTP(αS), 2′-deoxythymidine5′-O-(1-thiotriphosphate); dsDNA, double-stranded DNA; ELISA, enzyme-linked immunosorbent assay; SPR, surface plasmon resonance; KD, dissociation constant; FBS, fetal bovine serum; RT-PCR, real-time reverse transcription PCR; vRAGE, v-domain of RAGE; QCM, quartz crystal microbalance. ⁎ Correspondence to: Y. Higashimoto, Department of Chemistry, Kurume University School of Medicine, 67 Asahi-machi, Kurume 830-0011, Japan. ⁎⁎ Correspondence to: S. Yamagishi, Department of Pathophysiology and Therapeutics of Diabetic Vascular Complications, Kurume University School of Medicine, 67 Asahi-machi, Kurume 830-0011, Japan. E-mail addresses: [email protected] (Y. Higashimoto), [email protected] (S. Yamagishi). 0026-2862/$ – see front matter © 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.mvr.2013.08.010
of occupational age (L'Esperance et al., 1990). Development of diabetic retinopathy is characterized by loss of pericytes, increased vascular permeability and acellular capillaries, followed by microvascular thrombus formation in the retinas (Orlidge and D'Amore, 1987; Pfister et al., 2010; Yamagishi and Imaizumi, 2005a). Diabetic retinopathy ultimately progresses to proliferative changes associated with neovascularization (Yamagishi et al., 1993). Non-enzymatic modification of proteins by reducing sugars, a process also known as the Maillard reaction, has progressed at an extremely accelerated rate under diabetes, leading to the formation of advanced glycation end products (AGEs) (Yamagishi and Imaizumi, 2005b). There is accumulating evidence that AGEs are implicated in the development and progression of many pathological sequelae of diabetes- and age-associated disorders (Barile and Schmidt, 2007; Rahbar and Figarola, 2003; Sourris and Forbes, 2009; Sun et al., 2011; Yamagishi and Imaizumi, 2005b; Zong et al., 2011), including diabetic retinopathy. Indeed, we have previously shown that AGEs not only inhibit DNA synthesis, but also induce apoptotic cell death in cultured retinal pericytes, the earliest histopathological hallmark of diabetic retinopathy (Yamagishi and Imaizumi, 2005a). Moreover, we have found that AGEs evoke vascular hyperpermeability and
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Library and polymerase chain reactions (PCR)
thrombogenic reactions in endothelial cells (ECs) by inducing vascular endothelial growth factor (VEGF) and plasminogen activator inhibitor1 (PAI-1) through the interaction with the receptor for AGEs (RAGE), thereby being involved in diabetic retinopathy (Matsui et al., 2010; Ojima et al., 2012, Yamagishi et al., 1998, 2006, 2008). These observations suggest that blockade of AGEs–RAGE axis in retinal vasculatures may be a novel therapeutic target for diabetic retinopathy. In 1990s, an in vitro-selection process called systematic evolution of ligands by exponential enrichment (SELEX) was developed to screen single stranded nucleic acid molecules that bind specific ligands from random pool of library (Ellington and Szostak, 1990; Tuerk and Gold, 1990). These classes of single stranded molecules are referred as “aptamers”. Aptamers have possessed a high binding affinity and specificity to target proteins, whose property is more than or equal to that of monoclonal antibodies. In addition, its small size, non-immunogenicity and ease of modification compared to conventional monoclonal antibodies make aptamers a more attractive tool for therapeutic application (Jayasena, 1999). One major problem that arises in the therapeutic application of aptamers is their instability; they are susceptible to enzymatic nuclease attack in the cellular and serum fluids (Agrawal et al., 1995). To circumvent this, aptamers are further chemically modified through addition of amino, fluoro, or O-methyl in the 2′-position of ribose sugar and/or conjugation to polyethylene glycol or cholesterol (Wang et al., 2011; Yang et al., 2011). However, these post-SELEX modification processes are time consuming and sometimes might lead to decrease the binding affinity of aptamers (Wang et al., 2011; Yang et al., 2011). Therefore, it is better to screen aptamers directly from modified oligonucleotide-containing library. Indeed, phosphorothioate aptamers have the advantages of their enhanced affinity, specificity, and higher stability due to the sulfur backbone modifications (Yang et al., 2006). In this study, we screened novel phosphorothioate aptamers directed against AGEs (AGEs–thioaptamers) using a combinatorial approach involving the construction and screening of a phosphorothioate DNA library, and investigated if AGEs–thioaptamers could actually block the binding of AGEs to RAGE. We further examined the effects of AGEs–thioaptamers on DNA synthesis in AGE-exposed bovine retinal pericytes and studied whether they also could inhibit the AGE-induced RAGE, VEGF and PAI-1 gene expression in human umbilical vein EC (HUVEC).
The 80-nucleotide PCR product was applied to 20 μl of a Streptavidin Mag Sepharose (GE Healthcare) bead matrix suspended in binding/ washing buffer (2 M NaCl, 1 mM ethylenediaminetetraacetic acid, 10 mM Tris–HCl, pH 7.5). After equilibration of binding of the biotinylated double-stranded DNA (dsDNA) to streptavidin beads, unbound dsDNA was removed with binding/washing buffer, and matrixbound dsDNA was denatured. Then ssDNA was mixed with AGEs–HSA beads, and bound ssDNA was isolated as described previously (Higashimoto et al., 2007). To remove the ssDNA that could bind to non-glycated HSA, isolated ssDNA was passed through non-glycated HSA-immobilized agarose beads. The recovered ssDNA was amplified by PCR and used as the input DNA for the next selection. The sequences of the ssDNA cloned after repeating the SELEX procedure fifteen times were determined as described previously (Higashimoto et al., 2007).
Materials and methods
Enzyme-linked immunosorbent assay (ELISA)
Materials D-glyceraldehyde was purchased from Nakalai Tesque (Kyoto, Japan), and [3H]thymidine from GE Healthcare (Buckinghamshire, UK). Other chemicals were purchased from Sigma (St Louis, MO, USA).
Each well was coated with polyclonal antibodies raised against AGEs overnight. The wells were incubated with 10 μg/ml AGEs–HSA for 30 min, and then with 5′-biotin-labeled 250 nM AGEs–thioaptamers (Tanaka et al., 2009). After 30 min, horseradish peroxidase-streptavidin was added, and absorbance at 450 nm was measured.
Preparation of AGE-modified proteins
Surface plasmon resonance (SPR)
AGE-modified proteins were prepared as described previously (Yamagishi et al., 2002a). In brief, human serum albumin (HSA) or bovine serum albumin (BSA) was incubated under sterile conditions with D-glyceraldehyde for 7 days. Then, unincorporated sugars were then removed by dialysis against phosphate-buffered saline. Control non-glycated HSA or BSA was incubated under the same conditions except for the absence of reducing sugars.
AGEs–HSA and non-glycated HSA were immobilized via the amino groups to CM5 sensor chip (GE Healthcare) with the aid of 1-ethyl-3(3-dimethylaminopropyl)-carbodiimide and N-hydroxysuccinimide, respectively. The association and dissociation phases were monitored in a BIAcore 1000 (GE Healthcare). Chemically synthesized AGEs– thioaptamers were injected into the flow cell at concentrations of 5 and 10 nM at a flow rate of 10 μl/min. The sensor chip was regenerated with pulses of 20 mM Tris–HCl buffer (pH 8.0) containing 6 M urea to the baseline level, followed by an extensive washing with the running buffer. Control experiments were performed with ligand-free channel on the same sensor chip. From the assay curves obtained, the control signals, reflecting the bulk effect of buffer, were subtracted using BIAevaluation 4.1 software (GE Healthcare). Equilibrium dissociation constant (KD) was determined using the equation for 1:1 Langmuir binding.
Immobilizing AGEs–HSA on agarose beads AGEs–HSA was covalently coupled via sulfhydryl groups to iodoacetyl groups on SulfoLink Coupling Gel (Pierce, Rockford, IL, USA) as described previously (Higashimoto et al., 2007).
A random combinatorial single-stranded DNA (ssDNA) library with normal phosphate ester backbone oligonucleotides (80-mer) was synthesized (Greiner Bio-One, Tokyo, Japan): 5′-AGCTCAGAATGGATCC AAAC-[N]40-CATGAGAATTCGGCCGGATC-3′ where N is a randomized nucleotide with equal proportion of A, G, C, and T. The library with phosphorothioate backbone substituted at A and T positions was then synthesized by PCR amplification of the template using Ex Taq polymerase (Takara Bio, Otsu, Japan) and a mixture of 2′-deoxyadenosine5′-O-(1-thiotriphosphate) (dATP(αS)), 2′-deoxythymidine-5′-O(1-thiotriphosphate) dTTP(αS) (BIOLOG Life Institute, Bremen, Germany), dGTP and dCTP. The PCR condition for amplification of the starting random library (440 sequences) includes 200 μM each of dATP (αS), dTTP(αS), dGTP, and dCTP, 4 mM MgCl2, 740 nM 80-mer random template, 50 units of Ex Taq polymerase, and 2.4 μM each primer in a total volume of 0.1 ml. PCR was performed to amplify with 5′ primer (5′-AGCTCAGAATGGATCCAAAC-3′) and biotin-conjugated 3′ primer (biotin-5′-GATCCGGCCGAATTCTCATG-3′) (Greiner Bio-One) under the following conditions: 95 °C for 2 min; 30 cycles at 95 °C for 1 min, 52 °C for 30 s, and 72 °C for 15 s.
Selection of AGEs–thioaptamers
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Table 1 The sequences in random region of the selected AGEs–thioaptamers.
Bovine retinal pericytes were isolated and maintained in Dulbecco's modified Eagle medium (Gibco BRL, Rockville, USA) supplemented with 20% fetal bovine serum (FBS) (ICN Biomedicals Inc., Aurora, Ohio, USA) as described previously (Capetandes and Gerritsen, 1990). Cells at 5 to 10 passages were used for experiments. AGE treatments were carried out in a medium containing 2% FBS. HUVECs were cultured in endothelial basal medium supplemented with 2 %FBS, 0.4% bovine brain extracts, 10 ng/ml human epidermal growth factor and 1 μg/ml hydrocortisone according to the supplier's instructions (Clonetics Corp., San Diego, CA, USA). AGE treatment was carried out in a medium lacking epidermal growth factor and hydrocortisone.
Aptamers
Sequences of random region
#1s #2s #3s #4s #5s #6s #7s #8s #9s #10s #11s
ccAcgcAcATTAgcAAcccccAgcTcAcAcccATgAcAAA gTTgccgAcAAATTgAgTccAAcATAcAcgggggAAcAAc TgATcTgAAcATggTTcgTAcAcgATccccTgccccgAAA TgTAgcccgAgTATcATTcTccATcgcccccAgATAcAAg cAgAATcggggAccAcgAcAcTgcAcATAccTcgTAcgAA cgAcgTcccgccTcgATAATcAAccAAcAgccccggcTAA TAcAgcccggcAAATATTcccccATcTTgccgccTTcAAA TTTgcgAcgTAggATccTcAcccAcccgcgAAcgTcAAAA cATAcgcgTccAcAcATcTAAccccccTcAcAcccgcAAA TcTgccAcccTccgAcTAAcATATccggccTgAgAccAAA cAgAAcgAAccAccgcccAcAcgcTAcAcTAccAAccAAA
A and T, 5′ side of which are phosphorothioates, are in capital letters. 3
Measurement of [ H]thymidine incorporation into pericytes Pericytes were incubated with 100 μg/ml AGEs–BSA or non-glycated BSA in the presence or absence of the indicated concentrations of AGEs– thioaptamers for 20 h. [3H]thymidine incorporation into pericytes was measured as described previously (Higashimoto et al., 2007). Real-time reverse-transcription PCR (RT-PCR) HUVECs were treated with 100 μg/ml AGEs–BSA or non-glycated BSA for 4 h in the presence or absence of 2 or 20 nM clone #4s AGEs– thioaptamer. Then total RNAs were extracted with RNA queous-4 PCR kit (Ambion Inc., Austin, TX, USA) according to the manufacturer's instructions. Quantitative real-time RT-PCR was performed using Assayon-Demand and TaqMan 5 fluorogenic nuclease chemistry (Applied Biosystems, Foster city, CA, USA) according to the supplier's recommendation. IDs of primers and probe for human RAGE, VEGF, PAI-1, and βactin gene were Hs00153957_m1, Hs00900055_m1, Hs01126606_m1, and Hs01060665_g1, respectively (Applied Biosystems, Foster city, CA, USA). Binding affinity of AGEs–HSA to RAGE The binding affinity of AGEs–HSA to extracellular AGE-binding v-domain of RAGE (vRAGE) was measured using sensitive 27-MHz quartz crystal microbalance (QCM) (Affinix Q; Initium, Tokyo, Japan) as described previously (Kaida et al., 2013). In brief, recombinant vRAGE was immobilized on a QCM surface. After adding AGEs–HSA to reaction vessel, the time course of the frequency decrease of bound vRAGE on the QCM was monitored. The binding affinity of AGEs–HSA to vRAGE was calculated from curve fitting to the QCM frequency decrease. Statistical analysis All values are presented as mean ± standard deviation. One-way analysis of variance followed Dunnett's test was performed for statistical comparisons; p-values of less than 0.05 were considered significant. Results In vitro-selection of AGEs–thioaptamers To obtain AGEs–thioaptamers, we performed the SELEX protocol with an aptamer library based on a random 40-nucleotide sequence. After the 15th rounds of selection, 24 clones were sequenced from the pool of selected single-stranded DNAs to obtain 11 unique sequences, indicating that some of the sequences among the 24 clones were identical and that multiple selection of the same clone occurred. The sequences of AGEs–thioaptamers are shown in Table 1.
Binding affinity of AGEs–thioaptamers to AGEs–HSA or AGEs–HSA to vRAGE We first studied whether AGEs–thioaptamers obtained above could bind to AGEs–HSA using a sandwich ELISA. As shown in Fig. 1, all the clones bound to AGEs–HSA. Among them, clones #4s, #7s and #9s aptamers had higher binding affinity than the others. Furthermore, SPR analysis confirmed that all the 3 clones bound to AGEs–HSA, but not non-glycated HSA; KD values of #4s, #7s and #9s were 0.63, 0.36, and 0.57 nM, respectively (Fig. 2). Structural analysis revealed that all of them had bulge–loop structures. When the sequences of 3 aptamers were compared, the clone #4s aptamer contained more consensus sequence (Fig. 3A). The secondary structure of #4s aptamer is shown in Fig. 3B. We next examined the effects of AGEs–thioaptamers on AGEs– vRAGE interaction. QCM is a technique to detect a mass change on an electrode at nanogram level from the resonance frequency change; when molecules bound on oscillating quartz crystal, oscillating frequency decreases in proportional to binding amount of molecules on the surface (Efremov et al., 2013). As shown in Fig. 4, the highest dose of 50 nM of #4s aptamer did not evoke any alteration in ΔF/Hz, whereas 0 nM did, suggesting that #4s aptamer dose-dependently inhibited the binding of AGEs–HSA to vRAGE immobilized on a QCM surface. Effects of AGEs–thioaptamers on [3H]thymidine incorporation into AGE-exposed pericytes We investigated whether AGEs–thioaptamers could inhibit the AGEinduced decrease in DNA synthesis in cultured retinal pericytes. As shown in Fig. 5, three clones (#4s, #7s and #9s) were found to dosedependently restore the decrease in DNA synthesis in AGE-exposed pericytes; 200 nM clones #4s and #7s aptamers significantly inhibited the AGE-induced decrease in DNA synthesis in pericytes. Effects of #4s aptamer on RAGE, VEGF, and PAI-1 gene expression in AGE-exposed HUVEC We examined the effects of clone #4s aptamer on RAGE, VEGF and PAI-1 gene expression in HUVEC. As shown in Fig. 6, AGEs significantly increased RAGE, VEGF and PAI-1 mRNA levels in HUVEC, all of which were completely blocked by the treatment with 20 nM clone #4s aptamer. Discussion Microvessels are composed of pericytes and endothelial cells (Yamagishi and Imaizumi, 2005b; Yamagishi and Matsui, 2011). We have previously shown that pericytes play an important role in the maintenance of microvascular homeostasis (Yamagishi and Imaizumi,
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Fig. 1. Comparison of binding affinity of each selected AGEs–thioaptamer using a sandwich ELISA. Each well in a 96-well plate was coated with polyclonal antibodies raised against AGEs overnight. Then 100 μL of AGEs–HSA (10 μg/ml) was added to each well. After 30 min, 5′-biotin-labeled AGEs–thioaptamers (100 μL; 250 nM) were added into each well. The absorbance of each sample was measured at 450 nm. n = 3.
2005b; Yamagishi and Matsui, 2011). Therefore, loss of pericytes, the earliest histopathological hallmark of diabetic retinopathy, could predispose the vessels to angiogenesis, thrombogenesis and EC injury, thus leading to full-blown clinical expression of diabetic retinopathy (Yamagishi and Imaizumi, 2005b; Yamagishi and Matsui, 2011). In this study, we found that AGEs–thioaptamers (#4s, #7s and #9s) inhibited the AGE-induced decrease in DNA synthesis in retinal pericytes. In addition, we found that these 3 aptamers tightly bound to AGEs–HSA with the KD value of 10−10 M and that #4s aptamer dose-dependently inhibited the binding of AGEs–HSA to vRAGE. AGEs have been shown to decrease DNA synthesis and induce apoptotic cell death of pericytes through the interaction with RAGE (Yamagishi and Imaizumi, 2005b; Yamagishi et al., 2002b). Given that AGE accumulation was increased in diabetic retinas (Endo et al., 2001; Kim et al., 2011; Stitt et al., 1997), our present study suggests that AGEs–
thioaptamers might protect against pericyte damage and loss in diabetic retinopathy by suppressing the harmful effects of AGEs via blockade of their interaction with RAGE. In this study, we also found that AGEs–thioaptamer (#4s aptamer) inhibited the AGE-induced RAGE, VEGF and PAI-1 gene expression in HUVEC. AGEs up-regulated RAGE mRNA levels in EC, which may transduce the AGE signals again, thus further exacerbating the deleterious effects of AGEs (Ishibashi et al., 2010, Ojima et al., 2012; Yamagishi et al., 1998, 2006, 2008). Therefore, AGEs–thioaptamer (#4s aptamer) could reduce RAGE gene expression in AGE-exposed HUVEC by blocking the AGEs–RAGE signaling. Furthermore, AGEs–RAGE interaction has been shown to up-regulate VEGF and PAI-1 mRNA levels in EC (Yamagishi et al., 2006, 2008). VEGF is a specific mitogen for EC, also known as vascular permeability factor, and is considered to be a pivotal factor in the pathogenesis of various stages of diabetic retinopathy, including retinal
Fig. 2. Binding sensorgrams of AGEs–thioaptamers to immobilized AGEs–HSA and non-glycated HSA. The clones #4s (dotted line), #7s (solid line) and #9s (dashed line) were injected onto the sensor chip immobilized AGEs–HSA (bold lines) or non-glycated HSA (thin lines) at concentration of 10 nM, respectively. n = 3.
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Fig. 3. Analysis of consensus nucleotide sequences of clones #4s, #7s and #9s aptamers. (A) The common consensus sequence was obtained using Clustal W algorithm. The bold letters are used to indicate the consensus sequences that are conserved in all of three aptamers, and the underline letters indicate the sequences conserved in two aptamers. N represents the non-consensus sequence. (B) The predicted structure of clone #4 aptamer.
vascular hyperpermeability and neovascularization (Yamagishi and Imaizumi, 2005b; Yamagishi and Matsui, 2011; Yamagishi et al., 2006). Moreover, although microthrombosis is not a primary event in the evolution of diabetic retinopathy and that vasoregression is followed by pericyte loss (Pfister et al., 2010), PAI-1 could facilitate microthrombosis
formation and contribute to capillary obliteration and retinal ischemia in the diabetic retinas, further augmenting the VEGF expression and exacerbating diabetic retinopathy (Brazionis et al., 2008; Yamagishi and Imaizumi, 2005b; Yamagishi and Matsui, 2011). Taken together, the present observations suggest that blockade by AGEs–thioaptamers of the AGEs–RAGE axis in both pericytes and EC might be a novel therapeutic strategy for the treatment of diabetic retinopathy. We have previously shown that non-modified DNA aptamer raised against AGEs could bind to AGEs and neutralize their harmful effects on pericytes (Higashimoto et al., 2007). However, the binding affinity of non-modified AGEs–aptamer was not so high, and its KD value was about 10−6 M. So, in this study, we screened AGEs–thioaptamers, because this type of aptamers has been shown to exhibit not only longer lifetime under various biological milieus, but also better binding affinity and specificity to their target proteins (Lin et al., 1996; Yang et al., 2006). Indeed, the KD value of AGEs–thioaptamers used here was 104 times lower than that of non-modified AGEs–aptamer previously reported (Higashimoto et al., 2007). The theoretical structure of the clone #4s aptamer revealed the typical bulge–loop structure with cytosine-rich sequences such as ACC(C) or (C)CCA, whose structure was similar to that of the previously identified non-modified AGEs– aptamers (Higashimoto et al., 2007). These findings suggest that AGEs–thioaptamers may be more stable with resistance to nuclease attack compared with non-modified aptamers. Bulge–loop structure with ACC(C) or (C)CCA motif might be necessary for exclusive recognition of, and binding to, AGE-modified proteins. In this study, in vitro-modified AGEs were prepared by incubating HSA with glyceraldehyde for 1 week; this process produces relatively highly-modified proteins in comparison to those in vivo. However, it is unlikely that extensively-modified, nonphysiologic AGEs that were formed under the in vitro-conditions may exert non-specific and toxic effects on pericytes for the following reasons: we have previously found that (1) immunological epitope of glyceraldehyde-modified AGEs is actually present in serum of diabetic patients, (2) concentration (100 μg/ml) of in vitro-prepared AGEs used here is comparable with that of the in vivo-diabetic situation, (3) AGE-rich serum fractions derived from diabetic patients on hemodialysis evoke neuronal cell death in vitro, which was completely blocked by antibodies raised
Fig. 4. Frequency response plots of the inhibitory effect of clone #4s aptamer on AGEs–vRAGE interaction. Recombinant human vRAGE was immobilized on a QCM surface. After adding AGEs–HSA to the reaction vessel in the presence or absence of varied concentration of clone #4s aptamer, the time course of the frequency decrease of bound AGEs–HSA on the QCM was monitored. Clone #4s aptamer: 0 nM (solid line), 10 nM (dashed line), 50 nM (dotted line). n = 3.
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Fig. 5. Effects of AGEs–thioaptamers on DNA synthesis in AGE-exposed pericytes. Pericytes were incubated with 100 μg/ml AGE–BSA or non-glycated BSA in the presence or absence of 2, 20 or 200 nM AGEs–thioaptamers (#4s, #7s and #9s) for 20 h. Then, [3H]thymidine was added to the medium. After 4 h, [3H]thymidine incorporation into the cells was determined. *p b 0.05 and **p b 0.01 compared with the values of AGEs–BSA alone. n = 8.
against glyceraldehyde-derived AGEs (Takeuchi et al., 2000a, 2000b), and (4) AGEs–thioaptamers actually bind to glyceraldehyde-modified AGEs–HSA in this experiments. In the present study, we used HUVEC, EC type of large artery origin. Although we have previously shown that the response to AGEs in HUVEC (VEGF, RAGE, PAI-1 expression) is almost equal to that in capillary EC (Yamagishi et al., 1998, 2008), it would helpful to examine the effects of AGEs and AGEs–thioaptamers on human retinal capillary EC.
Acknowledgments This work was supported in part by MEXT-Supported Program for the Strategic Research Foundation at Private Universities, the Ministry of Education, Culture, Sports, Science and Technology (MEXT) (SY). This work was supported in part by Grants-in-Aid for Scientific Research (C) 25440056 (YH) and for Scientific Research (B) 22390111 (SY) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan. There is no conflict of the interest in this paper.
Fig. 6. Effect of clone #4s aptamer on (A) RAGE, (B) VEGF, and (C) PAI-1 gene expression in HUVEC. HUVECs were treated with 100 μg/ml AGEs–BSA or non-glycated BSA for 4 h in the presence or absence of 2 or 20 nM clone #4s aptamer. Total RNAs were transcribed and amplified by real-time PCR. Data were normalized by the intensity of GAPDH mRNA-derived signals and then related to the value obtained with non-glycated BSA. *p b 0.05 and **p b 0.01 compared with the values of AGEs–BSA alone. #p b 0.05 compared with the values of non-glycated BSA alone. n = 4.
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Sourris, K.C., Forbes, J.M., 2009. Interactions between advanced glycation end-products (AGE) and their receptors in the development and progression of diabetic nephropathy — are these receptors valid therapeutic targets. Curr. Drug Targets 10, 42–50. Stitt, A.W., Li, Y.M., Gardiner, T.A., et al., 1997. Advanced glycation end products (AGEs) co-localize with AGE receptors in the retinal vasculature of diabetic and of AGEinfused rats. Am. J. Pathol. 150, 523–531. Sun, J.K., Keenan, H.A., Cavallerano, J.D., et al., 2011. Protection from retinopathy and other complications in patients with type 1 diabetes of extreme duration: the Joslin 50-year medalist study. Diabetes Care 34, 968–974. Takeuchi, M., Makita, Z., Bucala, R., et al., 2000a. Immunological evidence that noncarboxymethyllysine advanced glycation end-products are produced from short chain sugars and dicarbonyl compounds in vivo. Mol. Med. 6, 114–125. Takeuchi, M., Bucala, R., Suzuki, T., et al., 2000b. Neurotoxicity of advanced glycation endproducts for cultured cortical neurons. J. Neuropathol. Exp. Neurol. 59, 1094–1105. Tanaka, Y., Honda, T., Matsuura, K., et al., 2009. In vitro selection and characterization of DNA aptamers specific for phospholamban. J. Pharmacol. Exp. Ther. 329, 57–63. Tuerk, C., Gold, L., 1990. Systematic evolution of ligands by exponential enrichment: RNA ligands to bacteriophage T4 DNA polymerase. Science 249, 505–510. Wang, R.E., Wu, H., Niu, Y., et al., 2011. Improving the stability of aptamers by chemical modification. Curr. Med. Chem. 18, 4126–4138. Yamagishi, S., Imaizumi, T., 2005a. Pericyte biology and diseases. Int. J. Tissue React. 27, 125–135. Yamagishi, S., Imaizumi, T., 2005b. Diabetic vascular complications: pathophysiology, biochemical basis and potential therapeutic strategy. Curr. Pharm. Des. 11, 2279–2299. Yamagishi, S., Matsui, T., 2011. Advanced glycation end products (AGEs), oxidative stress and diabetic retinopathy. Curr. Pharm. Biotechnol. 12, 362–368. Yamagishi, S., Kobayashi, K., Yamamoto, H., 1993. Vascular pericytes not only regulate growth, but also preserve prostacyclin-producing ability and protect against lipid peroxide-induced injury of co-cultured endothelial cells. Biochem. Biophys. Res. Commun. 190, 418–425. Yamagishi, S., Fujimori, H., Yonekura, H., et al., 1998. Advanced glycation endproducts inhibit prostacyclin production and induce plasminogen activator inhibitor-1 in human microvascular endothelial cells. Diabetologia 41, 1435–1441. Yamagishi, S., Inagaki, Y., Okamoto, T., et al., 2002a. Advanced glycation end productinduced apoptosis and overexpression of vascular endothelial growth factor and monocyte chemoattractant protein-1 in human-cultured mesangial cells. J. Biol. Chem. 277, 20309–20315. Yamagishi, S., Amano, S., Inagaki, Y., 2002b. Advanced glycation end products-induced apoptosis and overexpression of vascular endothelial growth factor in bovine retinal pericytes. Biochem. Biophys. Res. Commun. 290, 973–978. Yamagishi, S., Nakamura, K., Matsui, T., et al., 2006. Pigment epithelium-derived factor inhibits advanced glycation end product-induced retinal vascular hyperpermeability by blocking reactive oxygen species-mediated vascular endothelial growth factor expression. J. Biol. Chem. 281, 20213–20220. Yamagishi, S., Matsui, T., Nakamura, K., et al., 2008. Olmesartan blocks advanced glycation end products (AGEs)-induced angiogenesis in vitro by suppressing receptor for AGEs (RAGE) expression. Microvasc. Res. 75, 130–134. Yang, X., Wang, H., Beasley, D.W., et al., 2006. Selection of thioaptamers for diagnostics and therapeutics. Ann. N. Y. Acad. Sci. 1082, 116–119. Yang, Y., Ren, X., Schluesener, H.J., et al., 2011. Aptamers: selection, modification and application to nervous system diseases. Curr. Med. Chem. 18, 4159–4168. Zong, H., Ward, M., Stitt, A.W., 2011. AGEs, RAGE, and diabetic retinopathy. Curr. Diab. Rep. 11, 244–252.
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Brief Communication
Blockade by phosphorothioate aptamers of advanced glycation end products-induced damage in cultured pericytes and endothelial cells Yuichiro Higashimoto a,⁎, Takanori Matsui b, Yuri Nishino b, Junichi Taira a, Hiroyoshi Inoue c, Masayoshi Takeuchi d, Sho-ichi Yamagishi b,⁎⁎ a
Department of Chemistry, Kurume University School of Medicine, Kurume, Japan Department of Pathophysiology and Therapeutics of Diabetic Vascular Complications, Kurume University School of Medicine, Kurume, Japan Department of Chemistry, Keio University School of Medicine, Kanagawa, Japan d Department of Advanced Medicine Medical Research Institute, Kanazawa Medical University, Ishikawa, Japan b c
a r t i c l e
i n f o
Article history: Accepted 27 August 2013 Available online 5 September 2013
a b s t r a c t Advanced glycation end products (AGEs) not only inhibit DNA synthesis of retinal pericytes, but also elicit vascular hyperpermeability, pathological angiogenesis, and thrombogenic reactions by inducing vascular endothelial growth factor (VEGF) and plasminogen activator inhibitor-1 (PAI-1) through the interaction with the receptor for AGEs (RAGE), thereby being involved in the pathogenesis of diabetic retinopathy. In this study, we screened novel phosphorothioate-modified aptamers directed against AGEs (AGEs–thioaptamers) using a combinatorial chemistry in vitro, and examined whether these aptamers could inhibit the AGE-induced damage in both retinal pericytes and human umbilical vein endothelial cells (HUVECs). We identified 11 AGEs–thioaptamers; among them, clones #4, #7s and #9s aptamers had higher binding affinity to AGEs–human serum albumin (HSA) than the others. Surface plasmon resonance analysis revealed that KD values of #4s, #7s and #9s were 0.63, 0.36, and 0.57 nM, respectively. Furthermore, these 3 clones dose-dependently restored the decrease in DNA synthesis in AGE-exposed pericytes. AGEs significantly increased RAGE, VEGF and PAI-1 mRNA levels in HUVEC, all of which were completely blocked by the treatment with 20 nM clone #4s aptamer. Quartz crystal microbalance analysis confirmed that #4s aptamer dose-dependently inhibited the binding of AGEs–HSA to RAGE. Our present study demonstrated that AGEs–thioaptamers could inhibit the harmful effects of AGEs in pericytes and HUVEC by suppressing the binding of AGEs to RAGE. Blockade by AGEs–thioaptamers of the AGEs–RAGE axis might be a novel therapeutic strategy for diabetic retinopathy. © 2013 Elsevier Inc. All rights reserved.
Introduction Diabetic retinopathy is one of the most miserable complications in diabetes and is a leading cause of acquired blindness among the people Abbreviations: AGEs, advanced glycation end products; EC, endothelial cells; VEGF, vascular endothelial growth factor; PAI-1, plasminogen activator inhibitor-1; RAGE, receptor for AGEs; SELEX, systematic evolution of ligands by exponential enrichment; AGEs–thioaptamers, phosphorothioate aptamers directed against AGEs; HUVECs, human umbilical vein endothelial cells; HSA, human serum albumin; BSA, bovine serum albumin; ssDNA, single-stranded DNA; PCR, polymerase chain reactions; dATP(αS), 2′-deoxyadenosine-5′-O-(1-thiotriphosphate); dTTP(αS), 2′-deoxythymidine5′-O-(1-thiotriphosphate); dsDNA, double-stranded DNA; ELISA, enzyme-linked immunosorbent assay; SPR, surface plasmon resonance; KD, dissociation constant; FBS, fetal bovine serum; RT-PCR, real-time reverse transcription PCR; vRAGE, v-domain of RAGE; QCM, quartz crystal microbalance. ⁎ Correspondence to: Y. Higashimoto, Department of Chemistry, Kurume University School of Medicine, 67 Asahi-machi, Kurume 830-0011, Japan. ⁎⁎ Correspondence to: S. Yamagishi, Department of Pathophysiology and Therapeutics of Diabetic Vascular Complications, Kurume University School of Medicine, 67 Asahi-machi, Kurume 830-0011, Japan. E-mail addresses: [email protected] (Y. Higashimoto), [email protected] (S. Yamagishi). 0026-2862/$ – see front matter © 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.mvr.2013.08.010
of occupational age (L'Esperance et al., 1990). Development of diabetic retinopathy is characterized by loss of pericytes, increased vascular permeability and acellular capillaries, followed by microvascular thrombus formation in the retinas (Orlidge and D'Amore, 1987; Pfister et al., 2010; Yamagishi and Imaizumi, 2005a). Diabetic retinopathy ultimately progresses to proliferative changes associated with neovascularization (Yamagishi et al., 1993). Non-enzymatic modification of proteins by reducing sugars, a process also known as the Maillard reaction, has progressed at an extremely accelerated rate under diabetes, leading to the formation of advanced glycation end products (AGEs) (Yamagishi and Imaizumi, 2005b). There is accumulating evidence that AGEs are implicated in the development and progression of many pathological sequelae of diabetes- and age-associated disorders (Barile and Schmidt, 2007; Rahbar and Figarola, 2003; Sourris and Forbes, 2009; Sun et al., 2011; Yamagishi and Imaizumi, 2005b; Zong et al., 2011), including diabetic retinopathy. Indeed, we have previously shown that AGEs not only inhibit DNA synthesis, but also induce apoptotic cell death in cultured retinal pericytes, the earliest histopathological hallmark of diabetic retinopathy (Yamagishi and Imaizumi, 2005a). Moreover, we have found that AGEs evoke vascular hyperpermeability and
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Library and polymerase chain reactions (PCR)
thrombogenic reactions in endothelial cells (ECs) by inducing vascular endothelial growth factor (VEGF) and plasminogen activator inhibitor1 (PAI-1) through the interaction with the receptor for AGEs (RAGE), thereby being involved in diabetic retinopathy (Matsui et al., 2010; Ojima et al., 2012, Yamagishi et al., 1998, 2006, 2008). These observations suggest that blockade of AGEs–RAGE axis in retinal vasculatures may be a novel therapeutic target for diabetic retinopathy. In 1990s, an in vitro-selection process called systematic evolution of ligands by exponential enrichment (SELEX) was developed to screen single stranded nucleic acid molecules that bind specific ligands from random pool of library (Ellington and Szostak, 1990; Tuerk and Gold, 1990). These classes of single stranded molecules are referred as “aptamers”. Aptamers have possessed a high binding affinity and specificity to target proteins, whose property is more than or equal to that of monoclonal antibodies. In addition, its small size, non-immunogenicity and ease of modification compared to conventional monoclonal antibodies make aptamers a more attractive tool for therapeutic application (Jayasena, 1999). One major problem that arises in the therapeutic application of aptamers is their instability; they are susceptible to enzymatic nuclease attack in the cellular and serum fluids (Agrawal et al., 1995). To circumvent this, aptamers are further chemically modified through addition of amino, fluoro, or O-methyl in the 2′-position of ribose sugar and/or conjugation to polyethylene glycol or cholesterol (Wang et al., 2011; Yang et al., 2011). However, these post-SELEX modification processes are time consuming and sometimes might lead to decrease the binding affinity of aptamers (Wang et al., 2011; Yang et al., 2011). Therefore, it is better to screen aptamers directly from modified oligonucleotide-containing library. Indeed, phosphorothioate aptamers have the advantages of their enhanced affinity, specificity, and higher stability due to the sulfur backbone modifications (Yang et al., 2006). In this study, we screened novel phosphorothioate aptamers directed against AGEs (AGEs–thioaptamers) using a combinatorial approach involving the construction and screening of a phosphorothioate DNA library, and investigated if AGEs–thioaptamers could actually block the binding of AGEs to RAGE. We further examined the effects of AGEs–thioaptamers on DNA synthesis in AGE-exposed bovine retinal pericytes and studied whether they also could inhibit the AGE-induced RAGE, VEGF and PAI-1 gene expression in human umbilical vein EC (HUVEC).
The 80-nucleotide PCR product was applied to 20 μl of a Streptavidin Mag Sepharose (GE Healthcare) bead matrix suspended in binding/ washing buffer (2 M NaCl, 1 mM ethylenediaminetetraacetic acid, 10 mM Tris–HCl, pH 7.5). After equilibration of binding of the biotinylated double-stranded DNA (dsDNA) to streptavidin beads, unbound dsDNA was removed with binding/washing buffer, and matrixbound dsDNA was denatured. Then ssDNA was mixed with AGEs–HSA beads, and bound ssDNA was isolated as described previously (Higashimoto et al., 2007). To remove the ssDNA that could bind to non-glycated HSA, isolated ssDNA was passed through non-glycated HSA-immobilized agarose beads. The recovered ssDNA was amplified by PCR and used as the input DNA for the next selection. The sequences of the ssDNA cloned after repeating the SELEX procedure fifteen times were determined as described previously (Higashimoto et al., 2007).
Materials and methods
Enzyme-linked immunosorbent assay (ELISA)
Materials D-glyceraldehyde was purchased from Nakalai Tesque (Kyoto, Japan), and [3H]thymidine from GE Healthcare (Buckinghamshire, UK). Other chemicals were purchased from Sigma (St Louis, MO, USA).
Each well was coated with polyclonal antibodies raised against AGEs overnight. The wells were incubated with 10 μg/ml AGEs–HSA for 30 min, and then with 5′-biotin-labeled 250 nM AGEs–thioaptamers (Tanaka et al., 2009). After 30 min, horseradish peroxidase-streptavidin was added, and absorbance at 450 nm was measured.
Preparation of AGE-modified proteins
Surface plasmon resonance (SPR)
AGE-modified proteins were prepared as described previously (Yamagishi et al., 2002a). In brief, human serum albumin (HSA) or bovine serum albumin (BSA) was incubated under sterile conditions with D-glyceraldehyde for 7 days. Then, unincorporated sugars were then removed by dialysis against phosphate-buffered saline. Control non-glycated HSA or BSA was incubated under the same conditions except for the absence of reducing sugars.
AGEs–HSA and non-glycated HSA were immobilized via the amino groups to CM5 sensor chip (GE Healthcare) with the aid of 1-ethyl-3(3-dimethylaminopropyl)-carbodiimide and N-hydroxysuccinimide, respectively. The association and dissociation phases were monitored in a BIAcore 1000 (GE Healthcare). Chemically synthesized AGEs– thioaptamers were injected into the flow cell at concentrations of 5 and 10 nM at a flow rate of 10 μl/min. The sensor chip was regenerated with pulses of 20 mM Tris–HCl buffer (pH 8.0) containing 6 M urea to the baseline level, followed by an extensive washing with the running buffer. Control experiments were performed with ligand-free channel on the same sensor chip. From the assay curves obtained, the control signals, reflecting the bulk effect of buffer, were subtracted using BIAevaluation 4.1 software (GE Healthcare). Equilibrium dissociation constant (KD) was determined using the equation for 1:1 Langmuir binding.
Immobilizing AGEs–HSA on agarose beads AGEs–HSA was covalently coupled via sulfhydryl groups to iodoacetyl groups on SulfoLink Coupling Gel (Pierce, Rockford, IL, USA) as described previously (Higashimoto et al., 2007).
A random combinatorial single-stranded DNA (ssDNA) library with normal phosphate ester backbone oligonucleotides (80-mer) was synthesized (Greiner Bio-One, Tokyo, Japan): 5′-AGCTCAGAATGGATCC AAAC-[N]40-CATGAGAATTCGGCCGGATC-3′ where N is a randomized nucleotide with equal proportion of A, G, C, and T. The library with phosphorothioate backbone substituted at A and T positions was then synthesized by PCR amplification of the template using Ex Taq polymerase (Takara Bio, Otsu, Japan) and a mixture of 2′-deoxyadenosine5′-O-(1-thiotriphosphate) (dATP(αS)), 2′-deoxythymidine-5′-O(1-thiotriphosphate) dTTP(αS) (BIOLOG Life Institute, Bremen, Germany), dGTP and dCTP. The PCR condition for amplification of the starting random library (440 sequences) includes 200 μM each of dATP (αS), dTTP(αS), dGTP, and dCTP, 4 mM MgCl2, 740 nM 80-mer random template, 50 units of Ex Taq polymerase, and 2.4 μM each primer in a total volume of 0.1 ml. PCR was performed to amplify with 5′ primer (5′-AGCTCAGAATGGATCCAAAC-3′) and biotin-conjugated 3′ primer (biotin-5′-GATCCGGCCGAATTCTCATG-3′) (Greiner Bio-One) under the following conditions: 95 °C for 2 min; 30 cycles at 95 °C for 1 min, 52 °C for 30 s, and 72 °C for 15 s.
Selection of AGEs–thioaptamers
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Cells
Table 1 The sequences in random region of the selected AGEs–thioaptamers.
Bovine retinal pericytes were isolated and maintained in Dulbecco's modified Eagle medium (Gibco BRL, Rockville, USA) supplemented with 20% fetal bovine serum (FBS) (ICN Biomedicals Inc., Aurora, Ohio, USA) as described previously (Capetandes and Gerritsen, 1990). Cells at 5 to 10 passages were used for experiments. AGE treatments were carried out in a medium containing 2% FBS. HUVECs were cultured in endothelial basal medium supplemented with 2 %FBS, 0.4% bovine brain extracts, 10 ng/ml human epidermal growth factor and 1 μg/ml hydrocortisone according to the supplier's instructions (Clonetics Corp., San Diego, CA, USA). AGE treatment was carried out in a medium lacking epidermal growth factor and hydrocortisone.
Aptamers
Sequences of random region
#1s #2s #3s #4s #5s #6s #7s #8s #9s #10s #11s
ccAcgcAcATTAgcAAcccccAgcTcAcAcccATgAcAAA gTTgccgAcAAATTgAgTccAAcATAcAcgggggAAcAAc TgATcTgAAcATggTTcgTAcAcgATccccTgccccgAAA TgTAgcccgAgTATcATTcTccATcgcccccAgATAcAAg cAgAATcggggAccAcgAcAcTgcAcATAccTcgTAcgAA cgAcgTcccgccTcgATAATcAAccAAcAgccccggcTAA TAcAgcccggcAAATATTcccccATcTTgccgccTTcAAA TTTgcgAcgTAggATccTcAcccAcccgcgAAcgTcAAAA cATAcgcgTccAcAcATcTAAccccccTcAcAcccgcAAA TcTgccAcccTccgAcTAAcATATccggccTgAgAccAAA cAgAAcgAAccAccgcccAcAcgcTAcAcTAccAAccAAA
A and T, 5′ side of which are phosphorothioates, are in capital letters. 3
Measurement of [ H]thymidine incorporation into pericytes Pericytes were incubated with 100 μg/ml AGEs–BSA or non-glycated BSA in the presence or absence of the indicated concentrations of AGEs– thioaptamers for 20 h. [3H]thymidine incorporation into pericytes was measured as described previously (Higashimoto et al., 2007). Real-time reverse-transcription PCR (RT-PCR) HUVECs were treated with 100 μg/ml AGEs–BSA or non-glycated BSA for 4 h in the presence or absence of 2 or 20 nM clone #4s AGEs– thioaptamer. Then total RNAs were extracted with RNA queous-4 PCR kit (Ambion Inc., Austin, TX, USA) according to the manufacturer's instructions. Quantitative real-time RT-PCR was performed using Assayon-Demand and TaqMan 5 fluorogenic nuclease chemistry (Applied Biosystems, Foster city, CA, USA) according to the supplier's recommendation. IDs of primers and probe for human RAGE, VEGF, PAI-1, and βactin gene were Hs00153957_m1, Hs00900055_m1, Hs01126606_m1, and Hs01060665_g1, respectively (Applied Biosystems, Foster city, CA, USA). Binding affinity of AGEs–HSA to RAGE The binding affinity of AGEs–HSA to extracellular AGE-binding v-domain of RAGE (vRAGE) was measured using sensitive 27-MHz quartz crystal microbalance (QCM) (Affinix Q; Initium, Tokyo, Japan) as described previously (Kaida et al., 2013). In brief, recombinant vRAGE was immobilized on a QCM surface. After adding AGEs–HSA to reaction vessel, the time course of the frequency decrease of bound vRAGE on the QCM was monitored. The binding affinity of AGEs–HSA to vRAGE was calculated from curve fitting to the QCM frequency decrease. Statistical analysis All values are presented as mean ± standard deviation. One-way analysis of variance followed Dunnett's test was performed for statistical comparisons; p-values of less than 0.05 were considered significant. Results In vitro-selection of AGEs–thioaptamers To obtain AGEs–thioaptamers, we performed the SELEX protocol with an aptamer library based on a random 40-nucleotide sequence. After the 15th rounds of selection, 24 clones were sequenced from the pool of selected single-stranded DNAs to obtain 11 unique sequences, indicating that some of the sequences among the 24 clones were identical and that multiple selection of the same clone occurred. The sequences of AGEs–thioaptamers are shown in Table 1.
Binding affinity of AGEs–thioaptamers to AGEs–HSA or AGEs–HSA to vRAGE We first studied whether AGEs–thioaptamers obtained above could bind to AGEs–HSA using a sandwich ELISA. As shown in Fig. 1, all the clones bound to AGEs–HSA. Among them, clones #4s, #7s and #9s aptamers had higher binding affinity than the others. Furthermore, SPR analysis confirmed that all the 3 clones bound to AGEs–HSA, but not non-glycated HSA; KD values of #4s, #7s and #9s were 0.63, 0.36, and 0.57 nM, respectively (Fig. 2). Structural analysis revealed that all of them had bulge–loop structures. When the sequences of 3 aptamers were compared, the clone #4s aptamer contained more consensus sequence (Fig. 3A). The secondary structure of #4s aptamer is shown in Fig. 3B. We next examined the effects of AGEs–thioaptamers on AGEs– vRAGE interaction. QCM is a technique to detect a mass change on an electrode at nanogram level from the resonance frequency change; when molecules bound on oscillating quartz crystal, oscillating frequency decreases in proportional to binding amount of molecules on the surface (Efremov et al., 2013). As shown in Fig. 4, the highest dose of 50 nM of #4s aptamer did not evoke any alteration in ΔF/Hz, whereas 0 nM did, suggesting that #4s aptamer dose-dependently inhibited the binding of AGEs–HSA to vRAGE immobilized on a QCM surface. Effects of AGEs–thioaptamers on [3H]thymidine incorporation into AGE-exposed pericytes We investigated whether AGEs–thioaptamers could inhibit the AGEinduced decrease in DNA synthesis in cultured retinal pericytes. As shown in Fig. 5, three clones (#4s, #7s and #9s) were found to dosedependently restore the decrease in DNA synthesis in AGE-exposed pericytes; 200 nM clones #4s and #7s aptamers significantly inhibited the AGE-induced decrease in DNA synthesis in pericytes. Effects of #4s aptamer on RAGE, VEGF, and PAI-1 gene expression in AGE-exposed HUVEC We examined the effects of clone #4s aptamer on RAGE, VEGF and PAI-1 gene expression in HUVEC. As shown in Fig. 6, AGEs significantly increased RAGE, VEGF and PAI-1 mRNA levels in HUVEC, all of which were completely blocked by the treatment with 20 nM clone #4s aptamer. Discussion Microvessels are composed of pericytes and endothelial cells (Yamagishi and Imaizumi, 2005b; Yamagishi and Matsui, 2011). We have previously shown that pericytes play an important role in the maintenance of microvascular homeostasis (Yamagishi and Imaizumi,
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Fig. 1. Comparison of binding affinity of each selected AGEs–thioaptamer using a sandwich ELISA. Each well in a 96-well plate was coated with polyclonal antibodies raised against AGEs overnight. Then 100 μL of AGEs–HSA (10 μg/ml) was added to each well. After 30 min, 5′-biotin-labeled AGEs–thioaptamers (100 μL; 250 nM) were added into each well. The absorbance of each sample was measured at 450 nm. n = 3.
2005b; Yamagishi and Matsui, 2011). Therefore, loss of pericytes, the earliest histopathological hallmark of diabetic retinopathy, could predispose the vessels to angiogenesis, thrombogenesis and EC injury, thus leading to full-blown clinical expression of diabetic retinopathy (Yamagishi and Imaizumi, 2005b; Yamagishi and Matsui, 2011). In this study, we found that AGEs–thioaptamers (#4s, #7s and #9s) inhibited the AGE-induced decrease in DNA synthesis in retinal pericytes. In addition, we found that these 3 aptamers tightly bound to AGEs–HSA with the KD value of 10−10 M and that #4s aptamer dose-dependently inhibited the binding of AGEs–HSA to vRAGE. AGEs have been shown to decrease DNA synthesis and induce apoptotic cell death of pericytes through the interaction with RAGE (Yamagishi and Imaizumi, 2005b; Yamagishi et al., 2002b). Given that AGE accumulation was increased in diabetic retinas (Endo et al., 2001; Kim et al., 2011; Stitt et al., 1997), our present study suggests that AGEs–
thioaptamers might protect against pericyte damage and loss in diabetic retinopathy by suppressing the harmful effects of AGEs via blockade of their interaction with RAGE. In this study, we also found that AGEs–thioaptamer (#4s aptamer) inhibited the AGE-induced RAGE, VEGF and PAI-1 gene expression in HUVEC. AGEs up-regulated RAGE mRNA levels in EC, which may transduce the AGE signals again, thus further exacerbating the deleterious effects of AGEs (Ishibashi et al., 2010, Ojima et al., 2012; Yamagishi et al., 1998, 2006, 2008). Therefore, AGEs–thioaptamer (#4s aptamer) could reduce RAGE gene expression in AGE-exposed HUVEC by blocking the AGEs–RAGE signaling. Furthermore, AGEs–RAGE interaction has been shown to up-regulate VEGF and PAI-1 mRNA levels in EC (Yamagishi et al., 2006, 2008). VEGF is a specific mitogen for EC, also known as vascular permeability factor, and is considered to be a pivotal factor in the pathogenesis of various stages of diabetic retinopathy, including retinal
Fig. 2. Binding sensorgrams of AGEs–thioaptamers to immobilized AGEs–HSA and non-glycated HSA. The clones #4s (dotted line), #7s (solid line) and #9s (dashed line) were injected onto the sensor chip immobilized AGEs–HSA (bold lines) or non-glycated HSA (thin lines) at concentration of 10 nM, respectively. n = 3.
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Fig. 3. Analysis of consensus nucleotide sequences of clones #4s, #7s and #9s aptamers. (A) The common consensus sequence was obtained using Clustal W algorithm. The bold letters are used to indicate the consensus sequences that are conserved in all of three aptamers, and the underline letters indicate the sequences conserved in two aptamers. N represents the non-consensus sequence. (B) The predicted structure of clone #4 aptamer.
vascular hyperpermeability and neovascularization (Yamagishi and Imaizumi, 2005b; Yamagishi and Matsui, 2011; Yamagishi et al., 2006). Moreover, although microthrombosis is not a primary event in the evolution of diabetic retinopathy and that vasoregression is followed by pericyte loss (Pfister et al., 2010), PAI-1 could facilitate microthrombosis
formation and contribute to capillary obliteration and retinal ischemia in the diabetic retinas, further augmenting the VEGF expression and exacerbating diabetic retinopathy (Brazionis et al., 2008; Yamagishi and Imaizumi, 2005b; Yamagishi and Matsui, 2011). Taken together, the present observations suggest that blockade by AGEs–thioaptamers of the AGEs–RAGE axis in both pericytes and EC might be a novel therapeutic strategy for the treatment of diabetic retinopathy. We have previously shown that non-modified DNA aptamer raised against AGEs could bind to AGEs and neutralize their harmful effects on pericytes (Higashimoto et al., 2007). However, the binding affinity of non-modified AGEs–aptamer was not so high, and its KD value was about 10−6 M. So, in this study, we screened AGEs–thioaptamers, because this type of aptamers has been shown to exhibit not only longer lifetime under various biological milieus, but also better binding affinity and specificity to their target proteins (Lin et al., 1996; Yang et al., 2006). Indeed, the KD value of AGEs–thioaptamers used here was 104 times lower than that of non-modified AGEs–aptamer previously reported (Higashimoto et al., 2007). The theoretical structure of the clone #4s aptamer revealed the typical bulge–loop structure with cytosine-rich sequences such as ACC(C) or (C)CCA, whose structure was similar to that of the previously identified non-modified AGEs– aptamers (Higashimoto et al., 2007). These findings suggest that AGEs–thioaptamers may be more stable with resistance to nuclease attack compared with non-modified aptamers. Bulge–loop structure with ACC(C) or (C)CCA motif might be necessary for exclusive recognition of, and binding to, AGE-modified proteins. In this study, in vitro-modified AGEs were prepared by incubating HSA with glyceraldehyde for 1 week; this process produces relatively highly-modified proteins in comparison to those in vivo. However, it is unlikely that extensively-modified, nonphysiologic AGEs that were formed under the in vitro-conditions may exert non-specific and toxic effects on pericytes for the following reasons: we have previously found that (1) immunological epitope of glyceraldehyde-modified AGEs is actually present in serum of diabetic patients, (2) concentration (100 μg/ml) of in vitro-prepared AGEs used here is comparable with that of the in vivo-diabetic situation, (3) AGE-rich serum fractions derived from diabetic patients on hemodialysis evoke neuronal cell death in vitro, which was completely blocked by antibodies raised
Fig. 4. Frequency response plots of the inhibitory effect of clone #4s aptamer on AGEs–vRAGE interaction. Recombinant human vRAGE was immobilized on a QCM surface. After adding AGEs–HSA to the reaction vessel in the presence or absence of varied concentration of clone #4s aptamer, the time course of the frequency decrease of bound AGEs–HSA on the QCM was monitored. Clone #4s aptamer: 0 nM (solid line), 10 nM (dashed line), 50 nM (dotted line). n = 3.
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Fig. 5. Effects of AGEs–thioaptamers on DNA synthesis in AGE-exposed pericytes. Pericytes were incubated with 100 μg/ml AGE–BSA or non-glycated BSA in the presence or absence of 2, 20 or 200 nM AGEs–thioaptamers (#4s, #7s and #9s) for 20 h. Then, [3H]thymidine was added to the medium. After 4 h, [3H]thymidine incorporation into the cells was determined. *p b 0.05 and **p b 0.01 compared with the values of AGEs–BSA alone. n = 8.
against glyceraldehyde-derived AGEs (Takeuchi et al., 2000a, 2000b), and (4) AGEs–thioaptamers actually bind to glyceraldehyde-modified AGEs–HSA in this experiments. In the present study, we used HUVEC, EC type of large artery origin. Although we have previously shown that the response to AGEs in HUVEC (VEGF, RAGE, PAI-1 expression) is almost equal to that in capillary EC (Yamagishi et al., 1998, 2008), it would helpful to examine the effects of AGEs and AGEs–thioaptamers on human retinal capillary EC.
Acknowledgments This work was supported in part by MEXT-Supported Program for the Strategic Research Foundation at Private Universities, the Ministry of Education, Culture, Sports, Science and Technology (MEXT) (SY). This work was supported in part by Grants-in-Aid for Scientific Research (C) 25440056 (YH) and for Scientific Research (B) 22390111 (SY) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan. There is no conflict of the interest in this paper.
Fig. 6. Effect of clone #4s aptamer on (A) RAGE, (B) VEGF, and (C) PAI-1 gene expression in HUVEC. HUVECs were treated with 100 μg/ml AGEs–BSA or non-glycated BSA for 4 h in the presence or absence of 2 or 20 nM clone #4s aptamer. Total RNAs were transcribed and amplified by real-time PCR. Data were normalized by the intensity of GAPDH mRNA-derived signals and then related to the value obtained with non-glycated BSA. *p b 0.05 and **p b 0.01 compared with the values of AGEs–BSA alone. #p b 0.05 compared with the values of non-glycated BSA alone. n = 4.
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