Role of MicroRNA-214–Targeting Phosphatase and Tensin Homolog in Advanced Glycation End Product-Induced Apoptosis Delay in Monocytes

Monocyte activation, adhesion, and transmigration are key events in the pathogenesis of atherosclerosis (1). Advanced glycation end products (AGEs), well-known proinflammatory compounds, augment monocyte-mediated inflammatory responses via the receptor for AGEs (RAGE) (2–5). Driven by hyperglycemia, oxidant stress, and inflammation, proteins and lipids undergo nonenzymatic glycation and oxidation, generating new species distinct from the backbone protein or lipid structure. These products of nonenzymatic glycation accumulate under diverse dysfunctional conditions, including diabetes, inflammation, renal failure, and aging (2). For instance, dialysis-related amyloidosis (DRA) is a progressive and incapacitating condition that affects patients with chronic renal failure (CRF) (6, 7). In DRA, a histological hallmark is a local inflammatory reaction to β₂-microglobulin (β₂m) amyloid deposits mediated by monocytes and macrophages. β₂m modified with AGEs (β₂m-AGEs), a major component of amyloid in DRA (8), can affect the apoptosis and phenotype of human monocytes and macrophages (7). Meanwhile, an AGE-mediated increased resistance of macrophages to apoptosis in early lesions is associated with increased plaque burden in atherosclerosis (9). In general, through interaction with RAGEs, AGEs were shown to play a crucial role in the pathogenesis of such diabetic vascular complications as acute stroke, myocardial infarction, and peripheral vascular disease (10–12). However, although several signal pathways and interacting molecules, such as NF-κB (3) and RAGE (4, 5), were demonstrated to be involved in AGE-mediated modulation, the mechanisms by which AGEs modulate monocyte function, and particularly the apoptotic process, are not well understood.

Recently, the discovery of a new class of RNA regulatory genes, known as microRNAs (miRNAs), has revealed a whole new layer of gene regulation in eukaryotes. These miRNAs are endogenous, noncoding RNAs of 19–24 nucleotides in length that play an important role in the negative regulation of gene expression. This is accomplished by base-pairing to complementary sites on target mRNAs, thus blocking translation or causing degradation of the target gene (13). By negatively regulating their target gene expression, miRNAs play a critical role in diverse biological and pathological processes, including developmental timing, apoptosis, proliferation, differentiation, carcinogenesis, and the immune response (14, 15). However, it remains to be elucidated whether miRNAs play a role in the AGE-induced functional changes of monocytes. Little information about the role of miRNAs in monocytes under pathophysiological conditions, such as hyperglycemia and oxidant stress, is available.

In the current study, we used an miRNA microarray and stem-loop, quantitative RT-PCR (qRT-PCR) to identify differentially expressed miRNAs in monocyte/macrophage-lineage THP-1 cells after treatment with AGEs. AGE-induced differential expression of miRNAs was further examined in peripheral monocytes isolated from CRF patients, as well as from healthy donors. To our knowledge, this study provides the first evidence that AGEs can induce the upregulation of miRNA-214 (miR-214) in monocytes, which, in turn, plays a critical role in modulating monocyte survival by targeting the phosphatase and tensin homolog (PTEN).

THP-1 cells (China Cell Culture Center, Shanghai, China) were cultured as previously described (16). Human mononuclear leukocytes were isolated from the peripheral blood of healthy volunteers or patients with renal failure, as previously documented (17). Briefly, monocytes were isolated using Ficoll and anti-CD14 magnetic beads, according to the manufacturer’s instructions (Miltenyi Biotec, Auburn, CA). Monocytes isolated using this method were >90% pure, with no polymorphonuclear lymphocyte contamination. All procedures related to the handling of human blood were approved by the Institutional Review Board of Nanjing University and Nanjing Medical University, and written informed consent was obtained from each blood donor. Monocytes (4 × 10⁶/ml) were cultured in RPMI 1640 (Life Technologies BRL, Gaithersburg, MD) supplemented with 20 mM l-glutamine and 25 mM HEPES buffer. The viability of the monocytes was determined by trypan blue staining. An inhibitory anti-RAGE Ab was used, as previously described (18).

AGE-modified proteins were prepared in vitro, as previously described (7, 17). Briefly, 20 mg/ml endotoxin-free BSA (Fraction V, sterile-filtered; Sigma, St. Louis, MO) or 3 mg/ml normal human β₂m (Sigma) was incubated with 300 mM d-glucose and 150 mM lysine in 100 mM phosphate buffer containing 100 U/ml penicillin and streptomycin for 6 wk at 37°C. Samples incubated under identical conditions, but in the absence of glucose, served as controls. After incubation, all samples were dialyzed against PBS (pH 7.4) to remove unincorporated sugars. AGE reagents were further sterile filtered to exclude endotoxin contamination (19). AGE modification was characterized and quantitated by fluorospectrometry. The AGE content of the products was 41.7 U/mg protein for β₂m-AGEs, 42.9 U/mg protein for BSA modified with AGEs (BSA-AGEs), and 0.75 U/mg and 0.81 U/mg protein for the β₂m and BSA controls, respectively. All samples contained <5 pg/ml endotoxin, as measured by the limulus amebocyte lysate assay (Sigma) (17).

Serum deprivation-induced cell apoptosis (7) was assessed using Annexin V-FITC labeling (20) and caspase-3 activity measurement (21). Freshly isolated monocytes and monocytes treated with BSA-AGEs or β₂m-AGEs (BSA or β₂m served as controls) were washed repeatedly with PBS (pH 7.4) and then resuspended in ice-cold binding buffer (10 mM HEPES [pH 7.4], 140 mM sodium chloride, and 2.5 mM calcium chloride). The percentage of apoptotic cells was quantified using the FITC Annexin-V Apoptosis Detection Kit I (BD Pharmingen, San Jose, CA). Counterstaining with propidium iodide (PI; red fluorescence) was performed to discriminate between early apoptotic cells and cells at the end stage of apoptosis. Briefly, cells were incubated for 10 min at room temperature with 0.5 μg/ml Annexin V-FITC and 2 μg/ml PI, followed by washing with binding buffer. FACS analysis (FACScan; Becton Dickinson, Bedford, MA) was carried out immediately using standard protocols. Apoptotic cells were defined as those exhibiting exclusively green fluorescent signals. Cell apoptosis was also confirmed by fluorescent TUNEL and Hoechst 33258 staining (Invitrogen, Carlsbad, CA).

Total RNA was extracted from THP-1 cells using TRIzol reagent (Invitrogen) (22, 23). RNA labeling and hybridization on miRNA microarray chips were conducted as previously described (24). Briefly, 50 μg total RNA, purified using the mirVANA miRNA Isolation Kit (Applied Biosystems, Foster City, CA), was labeled with fluorescein, and hybridization was carried out on miRNA microarray chips (miRNA microarray V4.0; CapitalBio, Beijing, China) containing 1320 probes in triplicate. These probes corresponded to 988 human, 627 mouse, and 350 rat miRNA genes designed based on the miRBase Release 12.0. Three independent RNA samples from THP-1 cells treated with BSA-AGEs or BSA were hybridized with miRNA microarrays separately. Hybridization intensity values from individual samples were filtered and normalized to per-chip mean values. We considered candidate miRNAs with a signal >500 as positive.

Stem-loop qRT-PCR assays using TaqMan miRNA probes (Applied Biosystems) to quantify mature miRNAs were performed, as previously reported (23, 25). Real-time PCR was performed using a TaqMan PCR kit and the Applied Biosystems 7300 Sequence Detection System. The reactions were incubated in a 96-well optical plate at 95°C for 10 min, followed by 40 cycles of 95°C for 15 s and 60°C for 1 min. All reactions, including no-template controls, were run in triplicate. After the reactions, the cycle threshold (C_T) data were determined using default threshold settings, and the mean C_T was determined from the duplicate PCRs. A comparative ΔC_T method (26) was used to compare each condition with controls, and values are expressed as 2^−△CT. The relative levels of miRNAs in cells and tissues were normalized to U6, a ubiquitously expressed small nuclear RNA. All primers used are listed in Supplemental Table II.

A luciferase reporter assay was performed, as previously described (22), to test the binding of miR-214 to its target gene PTEN. A 1501-bp segment of human PTEN 3′-untranslated region (UTR), containing a presumed miR-214 complementary site (seed sequence, CCTGCTG), was amplified by PCR using human genomic DNA as a template. The PCR products were inserted into the pMIR-REPORT plasmid (Applied Biosystems), and efficient insertion was confirmed by sequencing. To test the binding specificity, we mutated the seed sequence of miR-214 from CCTGCTG to GGACGAC (Fig. 4A, inset). For the luciferase reporter assays, 1 μg firefly luciferase reporter plasmid, 1 μg β-galactosidase (β-gal) expression vector (Applied Biosystems), and 100 pmol pre–miR-214, miR-214 antisense oligonucleotide (ASO), or scrambled control RNA were transfected into 293T cells cultured in six-well plates using Lipofectamine 2000 (Invitrogen), according to the manufacturer’s instruction. The β-gal vector was used as a transfection control. One day after transfection, the cells were assayed using a luciferase assay kit (Promega, Madison, WI). The primers for miR-214 qRT-PCR assay were sense: 5′-GATGAGCTCAACTGAAGTGGCTAAAGAG-3′ and antisense: 5′-GATACGCGTTGAAGTTCTGCCTAATCTA-3′.

THP-1 cells (∼10⁶), seeded on six-well plates or 60-mm dishes, were transfected with pre–miR-214 or miR-214 ASOs using Lipofectamine 2000 (Invitrogen), according to the manufacturer’s instructions (22). For miR-214 overexpression, 100 pmol pre–miR-214 or scrambled control oligonucleotide was used. For knockdown of miR-214, 100 pmol miR-214 ASO or scrambled control oligonucleotide was used. Cells were harvested at 24 h posttransfection for semiquantitative RT-PCR and Western blotting analysis. For each well, an equal dose of miRNA precursors, ASOs, or scrambled control oligonucleotides was used.

The levels of PTEN, Akt, and phosphorylated Akt (pAkt) were analyzed by Western blots using polyclonal anti-human PTEN, polyclonal anti-Akt, and polyclonal anti-Thr(P)-308-Akt Abs (Cell Signaling, Danvers, MA), respectively. Normalization was performed by blotting the same samples with an Ab to β-actin.

All images of Western blots and qRT-PCR assays were representative of at least three independent experiments. Quantified microarray signal-intensity values were normalized to per-chip mean values. Differentially expressed miRNAs were identified using the t test procedure within significance analysis of microarrays (SAMs) (27), with a 5% false discovery rate threshold. Real-time PCR and luciferase reporter assays were performed in triplicate. Data shown are the mean ± SE for three or more independent experiments. Differences were considered statistically significant at p < 0.05, assessed using the Student t test (for paired samples) or Wilcoxon rank-sum (Mann–Whitney) test (for nonpaired samples).

Previous studies demonstrated that monocytes and macrophages express RAGEs and undergo activation and/or differentiation upon stimulation by AGEs (3). To test the hypothesis that AGE treatment maintains and aggravates inflammation by delaying the process of monocyte apoptosis and clearance, we determined the effects of AGEs on serum deprivation-induced apoptosis of monocytes by Annexin V binding and TUNEL assays. THP-1 cells, monocytes from healthy donors, and monocytes from patients with CRF were used in this study. The THP-1 cells and normal monocytes were treated with 100 μg/ml BSA-AGEs or β₂m-AGEs for various times, with cells treated with β₂m or BSA alone serving as controls. Because the plasma level of various AGEs, particularly β₂m-AGEs, is significantly higher in CRF patients than in healthy donors (28, 29), we regarded monocytes from these patients as AGE-treated monocytes and directly compared them with the monocytes isolated from healthy donors. As shown in Fig. 1, THP-1 cells treated with 100 μg/ml BSA-AGEs (Fig. 1C) had significantly delayed apoptosis after serum deprivation compared with BSA-treated THP-1 cells (Fig. 1B). As a control, THP-1 cells cultured in the presence of serum showed little apoptosis (Fig. 1A). After 48 h of incubation, nearly 60% of the BSA-treated THP-1 cells were apoptotic (25.7% late apoptosis, indicated by Annexin V and PI labeling; 34.3% early apoptosis, indicated by Annexin V-FITC labeling), whereas the apoptotic rate of THP-1 cells treated with BSA-AGEs was <40% (11.4% late apoptosis and 27.1% early apoptosis). Compared with the dynamics of THP-1 cell apoptosis in the absence of serum (Fig. 1D), apoptosis after AGE treatment was significantly slowed. AGEs had a similar effect on apoptosis in human monocytes isolated from healthy donors (Fig. 1E). At 48 h of incubation with serum deprivation, normal human monocytes treated with BSA-AGEs showed only half of the apoptotic rate of BSA-treated monocytes or nontreated monocytes. Reduced apoptosis of monocytes by BSA-AGEs was also confirmed by caspase-3 assay (Fig. 1F). As can be seen, monocytes treated with BSA-AGEs showed reduced levels of cleaved 19-kDa caspase-3 compared with BSA-treated monocytes, whereas the levels of procaspase-3 were not altered. Interestingly, when we compared normal human monocytes with those isolated from patients with CRF, who generally had a significantly greater concentration of β₂m-AGEs and other AGEs in plasma, we found that the monocytes from CRF patients exhibited a reduced rate of apoptosis (Fig. 1G). Moreover, in the absence of serum, cultured monocytes from CRF patients showed greater adhesion to the culture plates than did normal monocytes. The TUNEL assay indicated that the majority of these adhesive cells were not apoptotic (data not shown). Monocytes from CRF patients were also morphologically different from normal monocytes. As can be seen in Fig. 1H and 1I, after a 3-d culture, nearly 10% of total monocytes from patients displayed macrophage- or dendritic cell-like protuberances compared with monocytes from healthy donors, which showed no protuberances. Thus, the monocytes isolated from CRF patients may have differentiated under serum-free culture conditions.

To investigate whether circulating AGEs modulate monocytic function by inducing differential expression of miRNAs, we treated human THP-1 cells with BSA-AGEs or BSA alone overnight, followed by a microarray-based miRNA analysis (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE24885, accession number GSE24885). For microarray assay, only miRNAs with a signal >500 were considered positive. As shown in Fig. 2A and Supplemental Fig. 1, the patterns of miRNA expression in BSA-AGE– or BSA-treated THP-1 cells were significantly different. Using two-class unpaired analysis within SAM (27), the miRNA expression levels in BSA-AGE–treated THP-1 cells were compared with those of THP-1 cells treated with BSA alone. In the test, a false discovery rate (q-value) <5% was selected. The expression of miRNAs was only considered significantly altered by AGE treatment when it fulfilled two criteria: mean fold change >2 and q-value <5%. As shown in Supplemental Table I, SAM analysis generated a list of 46 miRNAs that were differentially expressed in THP-1 cells treated with BSA-AGEs compared with BSA-treated cells. Among these miRNAs, 41 were upregulated, whereas 5 were downregulated.

Next, we validated these differentially expressed miRNAs identified by the SAM algorithm based on the microarray results using TaqMan probe-based real-time qRT-PCR assay (25). We initially assessed most of these differentially expressed miRNAs (∼15% of total number of miRNA species) detected by microarray using qRT-PCR; however, we found, for some miRNAs with low copy numbers, substantial differences between the results derived from qRT-PCR and microarray analysis. Thus, we only selected the miRNAs with copy numbers >3000 in microarray detection from BSA-AGE–treated or BSA-treated THP-1 cells. Seven miRNAs, including miR-21, miR-99a, miR-122, miR-142-3p, miR-150, miR-155, and miR-214, were selected to be validated by qRT-PCR using RNA preparations isolated in six independent experiments. These miRNAs were selected because they are altered by AGE treatment (fold change >2), and they have a relatively high expression in THP-1 cells. As shown in Fig. 2B, miR-214, miR-150, and miR-99a were strongly upregulated by treatment with BSA-AGEs or β₂m-AGEs. In contrast, miR-21, miR-122, miR-142-3p, and miR-155 were not significantly altered. Because miR-21 was reported to be involved in modulating cell apoptosis, and its expression is altered in tumorigenesis (30–32), we further tested the levels of miR-21 and miR-214 in normal colon and colon cancer tissue (Fig. 2C). The results clearly showed an increase in miR-21 and miR-214 in colon cancer compared with normal colon tissue, which is in agreement with the previous finding (32, 33). This result validates the reliability of our assays in determining the expression levels of miR-21 and other miRNAs.

We then examined the role of BSA-AGEs and β₂m-AGEs in modulating the differential expressions of miRNAs using isolated human peripheral monocytes. In this experiment, freshly isolated monocytes from healthy donors were treated with BSA-AGEs or β₂m-AGEs for 12 h. Cells treated with BSA or β₂m alone served as controls. As shown in Fig. 3A, the expression of miR-214 was strongly upregulated by BSA-AGEs and β₂m-AGEs. The results confirmed that BSA-AGEs and β₂m-AGEs may specifically induce miR-214 expression in monocytes. Fig. 3B showed the dynamic change in miR-21, miR-150, and miR-214 in monocytes during AGE treatment. miR-214 was enhanced by AGEs after 12 h and was maintained at a high level during the course of treatment, whereas miR-21 was not changed. Interestingly, miR-150 was upregulated at 12 and 24 h of incubation with AGEs, but it decreased to almost the original level after 48 h of treatment. To elucidate whether AGE-induced monocytic miR-214 expression is involved in monocyte dysfunction during the pathophysiological process of CRF, we directly compared the miRNA-expression pattern of monocytes from CRF patients with that of normal human monocytes. The expression levels of these seven miRNAs in monocytes from 21 patients with CRF and 19 healthy donors were assessed by real-time qRT-PCR. As shown in Fig. 3C, the miR-214 levels in monocytes from CRF patients were >140-fold greater than those in control monocytes. This finding indicated that AGE-induced upregulation of miR-214 expression may serve as a molecular signature of chronic inflammatory conditions, such as in CRF. It is possible that upregulation of miR-214 by AGEs plays a critical role in modulating the inflammatory responses of monocytes under such pathophysiological conditions.

Next, we tested whether the effect of AGEs on upregulation of monocytic miR-214 expression is through binding to RAGE (3). In the experiment, isolated normal human monocytes were incubated with AGEs in the presence of 25 μg/ml inhibitory Ab against the extracellular domain of RAGE (anti-RAGE) or control Ab. As shown in Fig. 3D, upregulation of monocytic miR-214 by BSA-AGEs or β₂m-AGEs was largely abolished by anti-RAGE Ab, suggesting that the effect of AGEs on monocytic miR-214 expression is mainly through the AGE–RAGE pathway.

Using three computer-aided algorithms, TargetScan, miRanda, and PicTar, we obtained a list of predicted target genes that are potentially regulated by miR-214. One of the target genes, PTEN, which was predicted by two of three algorithms and was shown to play a critical role in cell survival in previous studies (34, 35), was selected for experimental verification. To test whether miR-214 targets PTEN, a 1501-bp segment of human PTEN 3′-UTR containing a possible miR-214 complementary site (Fig. 4A) was fused into a luciferase reporter plasmid. The resultant plasmid and control plasmid (β-gal) were introduced into 293T cells (22). Because miR-21 was also widely reported to target the PTEN (36–40), miR-19a (37), we further compared the PTEN-binding seed sequence between miR-21 and miR-214 (Fig. 4A). Fig. 4B shows the relative expression levels of miR-214 in 293T cells with and without transfection of scrambled precursor oligonucleotide (scramble), pre–miR-214, or miR-214 ASO (anti–miR-214). As can be seen, cellular miR-214 level was increased nearly 100-fold by overexpression of pre–miR-214. After being normalized to β-gal activity, luciferase reporter activity in 293T cells was decreased ∼40% by overexpression of pre–miR-214 (Fig. 4C) compared with the overexpression of scrambled precursor oligonucleotide. The reduction in luciferase reporter activity by overexpression of pre–miR-214 was abolished by cotransfection with anti–miR-214. Meanwhile, transfection of anti–miR-214 alone resulted in an ∼20% increase in luciferase reporter activity compared with the scrambled oligonucleotide-transfected cells. After mutating the nucleotides of seeding sequence in the PTEN 3′-UTR (Fig. 4A, inset), the inhibitory effect of pre–miR-214 and the promoting effect of anti–miR-214 on luciferase reporter activity were largely abolished (Fig. 4C). The results of the luciferase reporter activity assay demonstrated a direct binding of miR-214 to the 3′-UTR of PTEN, which may serve as a novel mechanism of miR-214–mediated PTEN-translation repression.

Next, we examined PTEN expression in THP-1 cells and monocytes treated with or without BSA-AGEs, as well as in monocytes isolated from CRF patients. As shown in Fig. 5, PTEN expression levels in THP-1 cells (Fig. 5A) and normal human monocytes (Fig. 5B) were significantly decreased after treatment with BSA-AGEs compared with cells treated with BSA alone. Because PTEN generally executes its role through downregulating Akt activity (41, 42), we further determined the level of pAkt in monocytes following AGE treatment (Fig. 5C). Compared with BSA-treated or nontreated monocytes, human normal monocytes treated with BSA-AGEs had an enhanced expression of pAkt, suggesting that the effect of AGEs on monocytic apoptosis delay is likely through the PTEN–Akt axis-signaling pathway. Fig. 5D shows the results of Western blotting analysis of monocytes isolated from 21 CRF patients and 19 healthy donors (some results, particularly from healthy donors, are not shown because of space limitations). Monocytes from CRF patients had significantly lower PTEN levels than did normal human monocytes. Semiquantitative analysis of monocytic PTEN levels in patients and healthy donors is shown in Fig. 5E. We also compared the pAkt level in monocytes from CRF patients and healthy donors; as shown in Fig. 5F, the pAkt level in monocytes from CRF patients was significantly higher than that in normal monocytes. We further measured the absolute level of monocytic miR-214 expression by constructing a standard curve using synthetic miR-214. As can be seen in Fig. 5G, the miR-214 level in monocytes from CRF patients was >40-fold higher than that in monocytes from healthy donors. The absolute level of miR-21, another miRNA that can target PTEN (36–40), was also assessed in the present experiments. In contrast, we found no significant increase in monocytic miR-21 levels in CRF patients compared with healthy donors, although the basal level of miR-21 was greater than miR-214 in monocytes from healthy donors.

Furthermore, we assessed PTEN levels in THP-1 cells after pre–miR-214 or anti–miR-214 transfection. As shown in Fig. 5H, overexpression of pre–miR-214 in THP-1 cells strongly increased cellular miR-214 expression but decreased PTEN levels in comparison with the control group. In contrast, compared with the control cells, overexpression of anti–miR-214 decreased cellular miR-214 expression but increased PTEN levels. The inverse correlation between miR-214 and PTEN expression levels in THP-1 cells or monocytes treated with AGEs, as well as monocytes from CRF patients, further supports the conclusion that monocytic PTEN is one of the target genes of miR-214.

Because it has been suggested that PTEN can modulate leukocyte apoptosis (34, 35), we postulated that miR-214–mediated targeting of PTEN may be a mechanism underlying the AGE-mediated delay in monocyte apoptosis. To test this hypothesis, gain-of-function and loss-of-function assays were used in THP-1 cells to investigate the functional roles of miR-214–targeting PTEN. As shown in Fig. 6, serum deprivation-induced apoptosis of THP-1 cells was significantly reduced by directly knocking down PTEN levels via PTEN small interfering RNA transfection. This finding indicated that PTEN is a candidate target gene linked to monocyte survival. An increase in miR-214 levels via overexpression of pre–miR-214 also strongly decreased the apoptosis of THP-1 cells, which is in agreement with our hypothesis that miR-214 targets PTEN and, thus, impedes THP-1 cell apoptosis. In contrast, downregulation of miR-214 in THP-1 cells via overexpression of anti–miR-214 increased the number of apoptotic cells. Furthermore, the effect of BSA-AGEs on serum deprivation-induced THP-1 cell apoptosis was largely abolished by transfecting THP-1 cells with anti–miR-214, implying that the role of BSA-AGEs in delaying monocyte apoptosis is through modulation of miR-214–targeting PTEN.

Monocytes are often described as orchestrators of the inflammatory response. For instance, recruitment of monocytes from the peripheral blood to the vascular intima is a central event in atherosclerosis (1). Although progress has been made in understanding the causes of monocyte activation, recruitment, and survival, the signaling events and genetic alterations involved remain unclear. Compelling evidence has demonstrated the regulatory role of miRNAs in monocytic functions. Consistent with these findings, we identified differentially expressed miRNAs induced by AGE stimulation of monocytes. These miRNAs are likely involved in AGE-induced monocyte survival during hyperglycemia.

Previous studies revealed dysregulated miRNA expression in monocytes under various inflammatory conditions (43, 44). However, although AGEs are major proinflammatory factors (5, 45), AGE-mediated alteration of monocytic miRNA expression has not been well characterized. Using an miRNA microarray, we surveyed the differential expression of miRNAs after AGE treatment. The results showed that a panel of miRNAs in AGE-treated THP-1 cells were significantly dysregulated (fold change >2) compared with those of THP-1 cells treated with BSA alone. To our knowledge, this is the first report demonstrating an alteration in monocytic miRNA expression at a genome-wide level due to treatment with AGE reagents. The distinctive miRNA-expression pattern induced in THP-1 cells by AGEs, and particularly the increase in miR-214 levels, was further confirmed in monocytes isolated from CRF patients and normal human monocytes treated with BSA-AGEs or β₂m-AGEs. Previous studies demonstrated that miR-150, as a specific miRNA in monocytes/macrophages, plays a critical role in modulating monocyte differentiation and inflammatory responses (46, 47). Microarray and qRT-PCR analysis indicated that miR-150 was also increased in THP-1 cells and monocytes following AGE treatment, implicating a potential role for miR-150 in AGE-mediated functional alteration in monocytes. However, the role of miR-150 was not characterized in the current study, mainly because miR-214 had a much greater increase in monocytes from CRF patients compared with monocytes from healthy donors. Together, these findings indicate that the differentially expressed miRNAs are involved in the inflammatory responses of monocytes under such pathophysiological conditions as hyperglycemia and CRF.

Although the fold change in miR-214 in BSA-AGE–treated THP-1 cells compared with BSA-treated cells was not the highest noted in the miRNA microarray, we selected miR-214 because it was consistently increased in THP-1 cells and human monocytes after treatment with AGE reagents. More importantly, among all of the miRNAs that we tested, miR-214 had the greatest increase in monocytes from CRF patients compared with monocytes from healthy donors. During the validation process of differentially expressed miRNAs by TaqMan probe-based qRT-PCR, some miRNAs with a larger fold change in response to BSA-AGEs versus BSA, including miR-10a, miR-486-3p, miR-511, and miR-654-5p, were discarded because of their low signal (copy number <3000). We have several pieces of evidence to support that miR-214 may target PTEN in monocytes. First, bioinformatics analysis implicated PTEN as one of the target genes of miR-214, with relatively high conservation among various species. Second, a luciferase reporter assay showed the direct binding of miR-214 to the 3′-UTR region of PTEN. Third, there was a clear inverse relationship between cellular miR-214 level and PTEN expression in monocytes and monocytic THP-1 cells. Our results are in agreement with the findings of Yang et al. (48), who showed that deregulated miR-214 in human ovarian cancer induces tumor cell survival and cisplatin resistance by targeting PTEN.

During the past few years, several reports showed the regulation of PTEN by various miRNAs. Other than miR-214, miR-21 (36–40), miR-19a (37), miR-26a (49), miR-106b and miR-17-92 families (50), miR-205 (51), miR-216/217 (52), miR-221/222 (53), miR-486 (54), and miR-494 (55) target PTEN. Targeting by multiple miRNAs without the common seeding sequence showed that PTEN is an efficient target of miRNAs in various physiological or pathophysiological processes. However, none of those potential PTEN-targeting miRNAs, with the exception of miR-214, was identified as an AGE treatment-sensitive miRNA by our selection criteria. They had low signal heat (<3000) or <2-fold change (AGEs versus control <2). Although miR-21 was widely found to reduce PTEN levels in various cell types (36–40) and was abundantly expressed in human monocytes (Fig. 5G), we detected no significant alteration of miR-21 in monocytes from CRF patients compared with those from healthy donors. These results suggested that miR-214 is a major monocytic miRNA that is responsible for modulating monocyte PTEN level in the presence of AGEs.

Previous studies had shown that prolonged monocyte survival can be mediated by endotoxins, such as LPS (23); thus, it raised a plausible question that delayed apoptosis of monocytes during AGE treatment might be due to the effect of endotoxins in AGE reagents. However, three factors excluded the possibility of contamination by endotoxin: 1) the effect of AGE reagents on monocyte apoptosis can be diminished by inhibitory Ab against RAGE, a receptor of AGEs (Fig. 3D), suggesting that prolonged monocyte survival is due to AGEs but not endotoxin; 2) serving as control, BSA or β₂m was incubated with the same phosphate buffer (without glucose) within the same time frame; if the incubation caused generation of endotoxins, the BSA or β₂m solution should contain a similar level of endotoxin, but neither had an effect on monocyte survival; and 3) after dialysis and filtration, the level of endotoxin in prepared AGE reagents was low (<5 pg/ml). Although the underlying mechanism remains unknown, AGE–RAGE interactions were shown to limit cell apoptosis and promote cell survival and differentiation (7, 56, 57). Several possible pathways, including a p53-dependent mitochondrial pathway, are suggested to regulate cell apoptosis as a result of AGE–RAGE interactions (57). In the current study, we demonstrated that AGEs can strongly enhance miR-214 expression in monocytes, which, in turn, impedes monocyte apoptosis via targeting of the proapoptotic molecule PTEN. As an inhibitor of Akt activation, PTEN was demonstrated to be involved in leukocyte inflammatory responses (52, 58, 59). Downregulation of PTEN would result in increased Akt activity (Fig. 5C), leading to monocyte survival. Koide et al. (59) found that PTEN overexpression can reduce neointima formation, possibly through inhibition of macrophage invasion and proinflammatory cytokine expression. Working with alveolar macrophages, Flaherty et al. (58) found that PTEN was decreased following monocyte-to-macrophage differentiation and that PTEN deficiency in alveolar macrophages led to the enhancement of Akt activity and prolonged macrophage survival. Reduction in PTEN levels by other miRNAs, such as miR-21, was also shown to promote tumor cell survival and growth (36, 60).

Although our data demonstrated that miR-214 is significantly elevated in monocytes after AGE treatment or under hyperglycemia, the molecular basis underlying this event is unclear. Lee et al. (61) suggested that Twist-1 could regulate the miR-199a/214 cluster during development of specific neural cell populations via an E-Box promoter element. Meanwhile, Vinciguerra et al. (62) showed that the promoter activity of miR-21, another PTEN-targeting miRNA, was increased by mammalian target of rapamycin/NF-κB activation. It would be interesting to determine whether an AGE executes its role via similar pathways.

In conclusion, through an extensive analysis of miRNA expression in AGE-treated human monocytes and monocytes under chronic inflammatory conditions, such as renal failure and hyperglycemia, the current study demonstrated a distinctive miRNA-expression pattern in AGE-treated monocytes and a novel role for miR-214–targeting PTEN in AGE-induced monocyte resistance to apoptosis. Identification of the miR-214–targeting PTEN pathway in the modulation of monocytic apoptosis provides a potential new therapeutic target in the treatment of inflammatory disease.

We thank Dr. Xi Chen for assistance with data analysis and figure preparation.

The authors have no financial conflicts of interest.