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Division of Nephrology, Nanfang Hospital, Southern Medical University, Guangzhou 510515, People’s Republic of China
Address all correspondence and requests for reprints to: Dr. Fan Fan Hou, Division of Nephrology, Nanfang Hospital, Southern Medical University, 1838 North Guangzhou Avenue, Guangzhou 510515, People’s Republic of China. E-mail: ffhou{at}public.guangzhou.gd.cn.
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Abstract |
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Introduction |
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Diabetic nephropathy is generally considered a nonimmune disease. However, aberrant inflammatory responses, such as macrophage infiltration and overexpression of monocyte chemoattractant protein-1 (MCP-1), have been demonstrated in diabetic kidney (2, 3, 4). Macrophage recruitment and the associated inflammatory processes have been found in early diabetic renal changes as well as in established diabetic nephropathy in both human and animal models and have been linked to glomerular and tubular damage and renal fibrosis (2, 3, 5, 6, 7). Although these reports support the notion that inflammatory processes are implicated in development and/or progression of diabetic nephropathy, the mechanisms involved in the pathogenesis of diabetic renal inflammation have not been completely understood. It remains unclear whether renal inflammation in diabetes is simply a response to hyperglycemia or to obesity (3, 8) or whether any other metabolic factor plays a role in driving or promoting renal inflammation.
Diabetes is associated with increased modification of proteins. A typical representation is the formation and accumulation of advanced glycation end products (AGEs), the products of nonenzymatic glycation/oxidation of proteins/lipids. AGEs induce activation of protein kinase C and increase expression of TGF-β1 and extracellular matrix (ECM) in glomerular endothelial and mesangial cells (1). In addition to AGEs, a family of oxidized protein compounds, termed advanced oxidation protein products (AOPPs), has emerged as a novel class of inflammatory mediators. AOPPs are the dityrosine-containing and cross-linking protein products formed during oxidative stress (9, 10). Plasma AOPPs are mainly carried by albumin in vivo (9). Accumulation of AOPPs was first found in patients undergoing dialysis (9) and subsequently demonstrated in diabetic patients (11, 12) and in subjects with obesity (13). Our recent studies have found that iv infusion of AOPPs-albumin significantly increases macrophage infiltration in the remnant kidneys from rats receiving five sixths nephrectomy (14). Increased macrophage influx was also demonstrated in aorta from the hypercholesterolemic rabbits challenged by AOPPs (15). Consistent with the observations, accumulation of AOPPs has been linked to chronic inflammation and monocyte activation in patients with chronic renal insufficiency (10). These data suggest that AOPPs might be potential inducers of cellular inflammation in vivo under certain pathophysiological circumstances.
The present study was conducted to test the hypothesis that AOPPs accumulation may promote inflammation in diabetic kidney. Our data indicate that chronic administration of AOPPs in streptozotocin (STZ)-induced diabetes resulted in significant increase of macrophage infiltration and overexpression of MCP-1 and TGF-β1 in kidney. Renal inflammation was associated with deteriorated structural and functional abnormalities of diabetic kidney, such as glomerular hypertrophy, ECM accumulation and albuminuria. Our results also show that the proinflammatory effects of AOPPs on diabetic kidney might be mainly through activation of renal nicotinamide adenine dinucleotide phosphate (NADPH) oxidase, because AOPPs-induced renal inflammation was associated with increased activity of renal NADPH oxidase and could be prevented by inhibition of the enzyme. These data provide new insight on the pathogenesis of renal inflammation in diabetes and suggest that AOPPs accumulation might be a therapeutic target, at least for prevention or delaying the progression of renal complication in diabetes.
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Materials and Methods |
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AOPPs content in the preparation, as determined by the absorbance of the reaction mixture of the samples and acetic acid (9, 14), was 53.3 ± 4.0 nmol/mg protein in AOPPs-RSA and 0.3 ± 0.03 nmol/mg protein in native RSA. AOPPs levels in plasma and homogenate of renal tissue were quantified as described previously (9, 14).
To further determine whether the AOPPs preparation contains AGEs, we measured the content of N
-(carboxylmethyl)lysine, pentosidine, and glycolaldehyde-pyridine in both AOPPs-RSA and native RSA as described (15, 16, 17). There was no significant difference in pentosidine concentration between AOPPs-RSA (0.75 ± 0.06 nmol/liter) and native RSA (0.80 ± 0.07 nmol/liter, P > 0.05). The content of N-(carboxylmethyl)lysine was comparable between the AOPPs-RSA (41.0 ± 8.2 ng/mg protein) and unmodified RSA (38.2 ± 8.0 ng/mg protein, P > 0.05). Glycolaldehyde-pyridine was undetectable in AOPPs-RSA by using ELISA.
Animal models
Experimental diabetes was induced in male Sprague Dawley rats (initial weight, 230
250 g, Southern Medical University Animal Experiment Center) by ip injection of STZ (Sigma; 50 mg/kg) after an overnight fast. Animals with nonfasting serum glucose levels of more than 20 mmol/liter 1 wk after injection of STZ were included in the study as diabetic models. Diabetic rats were randomized into five groups (n = 12 in each group) matched for body weight and blood glucose and received the following treatment: group 1, iv injection of vehicle (PBS, pH 7.4); group 2, iv injection of native RSA at 50 mg/kg·d; group 3, iv injection of AOPPs-RSA at 50 mg/kg·d; group 4, iv injection of AOPPs-RSA at 50 mg/kg·d and apocynin (Sigma) at 100 mg/kg·d (18) in drinking water changed daily to quantify the volume intake; and group 5, apocynin at 100 mg/kg·d in drinking water. All the treatments were given from randomization until the animals were killed at wk 5 or 10 (n = 6 at each time point). The dosage of AOPPs and the injection interval were based on our preliminary studies, indicating that by this procedure, plasma AOPPs concentration increased by approximately 1-fold (the level that has been found in patients with diabetes) (11) in diabetic rats. Vehicle-injected (sodium citrate buffer, pH 4.5) control rats (group 6) were followed concurrently.
At the end of 5 and 10 wk after treatments, the animals (n = 6 in each group at each time point) were anesthetized with sodium pentobarbital and exsanguinated. The kidneys were collected after perfusion with 50 ml ice-cold normal saline. The 24-h urine samples were collected in metabolic cages at the end of the study period. Mean systolic blood pressure by tail cuff plethysmography (19) was measured before starting the treatment and at the end of the study period. All animal procedures were in accordance with guidelines set by the Animal Experiment Committee of Southern Medical University.
Renal morphometric analysis
Renal morphometric analysis was performed as described previously (20). A transversal section at the level of the hilus was fixed in periodate-lysine-paraformaldehyde solution and embedded in paraffin. Sections (2 µm thick) of the renal tissues were stained with periodic acid-Schiff for morphometric analysis. The measured glomerular parameters included 1) total glomerular surface area (Ag) and 2) mesangial surface area/glomerular tuft surface area. In each renal section, the observer blindly measured at least 30 consecutive unselected glomeruli randomly distributed over the depth of the cortex. Polar glomerular sections were not measured. Individual glomerular volume (Vg) was calculated as Vg = 1.25 x (Ag)3/2 (14).
Renal immunohistochemical analysis
The immunoperoxidase staining was performed as described (21), with the following antibodies as the primary antibodies: macrophage infiltration was detected with monoclonal anti-ED-1 (Serotec, Oxford, UK), TGF-β1 expression was detected with polyclonal rabbit antirat TGF-β1 (Santa Cruz Biotechnology, Santa Cruz, CA; catalog no. sc-146), MCP-1 was analyzed with polyclonal rabbit antirat MCP-1 (Boster Biologic, Wuhan, China; catalog no. BA1255), and fibronectin expression was analyzed with monoclonal antirat fibronectin (Labvision Corp., Fremont, CA). Control experiments included omission of the primary antibodies and substitution of the primary antibodies with nonimmune rabbit or mouse IgG.
Macrophage infiltration was quantified by counting the number of ED-1-positive cells in 30 glomerular profiles and in 20 randomly chosen 0.3 x 0.3-mm areas of tubulointerstitium for each kidney (14). The intensity of glomerular staining of TGF-β1, MCP-1, and fibronectin was evaluated under x400 magnification according to the following scale: 0, no staining; 1, weak and spotty intraglomerular staining; 2, moderate and segmental intraglomerular staining; and 3, strong and diffuse (involving >50%) intraglomerular staining. Likewise, the intensity of tubulointerstitial staining in cortical areas was assessed under x250 magnification according to the following scale: 0, no staining; 1, less than 25% involvement; 2, 25–50% involvement; and 3, more than 50% involvement. All but at least 20 glomeruli and 20 randomly selected cortical tubulointerstitial areas from each sample were evaluated.
RT-PCR
Total RNA was extracted from renal cortex using TRIZOL reagent (Invitrogen, Carlsbad, CA) according to the manufacturer’s protocol. Primers for MCP-1, TGF-β1, and glyceraldehyde-3-phosphate dehydrogenase were designed and synthesized on the basis of the published sequences of these genes (21). The upstream and downstream sequences of these primers are as follows: 1) MCP-1, 5'-GTC ACC AAG CTC AAG AGA GAG A-3' and 5'-GAG TGG ATG CAT TAG CTT CAG A-3'; 2) TGF-β1, 5'-CTT CAG CTC CAC AGA GAA GAA CTG C-3' and 5'-CAC GAT CAT GTT GGA CAA CTG CTC C-3'; and 3) glyceraldehyde-3-phosphate dehydrogenase, 5'-AAT GCA TCC TGC ACC ACC AA-3' and 5'-GTA GCC ATA TTC ATT GTC ATA-3'. RT-PCR was performed using TaKaRa RNA PCR kit version 3.0 (TaKaRa Biotechnology, Dalian, China) according to the manufacturer’s instructions. Target sequences were amplified at 56 C using the same amount of cDNA for all primer sets, and the cycle number was adjusted between 30 and 35 to yield visible products within the linear amplification range. RT-PCR products were resolved in 2% agarose gel and photographed under UV light. Density of bands was measured by scanning densitometry with UVIBAND V.99 software (UVI; SJ, Cambridge, UK).
Renal function and urine examination
Serum and urine creatinine levels were determined using commercial kits (sarcosine oxidase-peroxidase-antiperoxidase; Zixing, Shanghai, China) according to the manufacturer’s instruction. The creatinine clearance (Ccr) was calculated as described and factored for body weight (14). Twenty-four-hour urinary albumin excretion was measured as described previously (22).
Urinary MCP-1 and TGF-β1 were quantified by ELISA using commercial kits (BioSource, Camarillo, CA) according to the manufacturer’s instructions.
Estimation of renal NADPH oxidase activity
NADPH-dependent O2– production by homogenates from renal cortex was assessed by lucigenin-enhanced chemiluminescence as described previously (23). Homogenates were subjected to low-speed centrifugation (1000 x g) for 10 min. The supernatant (100 µg protein) was then added into a 96-well microplate containing 5 µmol/liter lucigenin and 100 µmol/liter NADPH. The chemiluminescence value was recorded at 30-sec intervals over 15 min (Victor β Wallac 1420; PerkinElmer, Turku, Finland), and readings in each of the last 5 min were averaged. Lucigenin count was expressed as counts/min·100 µg protein. The background was determined by measurement in the absence of homogenate and then subtracted from the reading obtained in the presence of homogenate. Generation of O2– was also determined in the presence of 10 µmol/liter diphenylene iodonium (DPI, an inhibitor of flavin-containing enzymes), 250 µmol/liter rotenone (a mitochondria inhibitor), 10 µmol/liter indomethacin (an inhibitor of cyclooxygenase), 10 µmol/liter NG-monomethyl-L-arginine (L-NMMA, an inhibitor of nitric oxide synthase), and 100 µmol/liter allopurinol (an inhibitor of xanthine oxidase) (all from Sigma).
Renal expression of NADPH oxidase subunits
Membrane proteins were extracted using a proteoExtract native membrane protein extraction kit (Calbiochem, San Diego, CA) according to the manufacturer’s instruction. Briefly, the kidney tissues were homogenized in ice-cold extraction buffer I containing protease inhibitor cocktail. Homogenates were centrifuged at 16,000 x g for 15 min at 4 C, and supernatant containing the cytosolic fraction was collected. The pellet was then incubated with ice-cold extraction buffer II supplemented with protease inhibitor cocktail for 30 min at 4 C under gentle agitation. The suspension was centrifuged at 16,000 x g for 15 min at 4 C to isolate the membrane fraction contained in the supernatant. Expression of NADPH oxidase subunits in the membrane and cytosolic fraction was analyzed by Western blot as described (24). Briefly, samples were boiled for 5 min. Aliquots (70 µg protein per lane for the membrane fraction, 30 µg protein per lane for the cytosolic fraction) were resolved with 15% SDS-polyacrylamide gels and transferred to nitrocellulose membranes (Amersham Pharmacia Biotech, Piscataway, NJ). The membranes were blocked with 5% nonfat milk in Tris-buffered saline containing 0.1% Tween 20, followed by overnight incubation with a rabbit antirat p47phox antibody (1:1000; Upstate, Lake Placid, NY), a mouse antirat gp91phox monoclonal antibody (1:500; BD Biosciences PharMingen, San Diego, CA), and a rabbit antirat p22phox antibody (1:200; Santa Cruz Biotechnology). After washing with Tris-buffered saline containing 0.1% Tween 20, the membranes were incubated with horseradish peroxidase-conjugated antirabbit or antimouse IgG second antibodies for 1 h at room temperature and detected by using ECL detection system (Pierce). The density of the bands was quantified by densitometry (Universal Hood.2; Bio-Rad, Milan, Italy). Blots were further probed with anti-actin antibody (Santa Cruz Biotechnology) to normalize for protein loading.
Statistical analysis
All data are expressed as mean ± SEM. Continuous variables between groups at each time point were compared using one-way ANOVA, followed by the least significant difference method when P 
0.05. Differences of the variables between two time points were determined by independent-samples t test. The relationship between variables was assessed by Pearson correlation analysis. Because of a positively skewed distribution, urinary albumin excretion was logarithmically transformed before statistical analysis and expressed as the geometric means x/÷ tolerance factor. Statistical analyses were conducted with SPSS 13.0 for Windows (SPSS, Chicago, IL). Significance was defined as P 0.05.
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Results |
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, diabetic rats showed significantly higher blood glucose levels, lower body weight, and greater kidney weight as compared with control rats. Mean systolic blood pressure was comparable among the groups.
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AOPPs administration increased AOPPs expression in diabetic kidney
Plasma AOPPs levels spontaneously increased in vehicle-treated diabetic rats compared with normal controls (P < 0.001, Fig. 1A). The concentration of plasma AOPPs increased by approximately 1-fold in AOPPs-RSA-treated diabetic rats at wk 10 compared with vehicle- or RSA-treated diabetic animals (Fig. 1A). In contrast, there was no significant difference in plasma albumin levels among groups at any time point (data not shown).
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). Chronic administration of AOPPs-RSA, but not native RSA, significantly enhanced AOPPs expression in diabetic kidney (Fig. 1B). Intervention of apocynin significantly decreased the renal AOPPs expression in both plasma and renal tissue (Fig. 1).
AOPPs administration promoted inflammation in diabetic kidney
Renal macrophage infiltration.
Few ED-1-positive macrophages could be found in renal tissues from normal control rats. As shown in Fig. 2, macrophage infiltration was evident in both glomeruli and interstitium of the diabetic kidney. At wk 5 after STZ injection, the number of infiltrated macrophages was increased markedly in AOPPs-RSA- vs. RSA- or vehicle-treated diabetic rats. These increases were sustained at wk 10 (Fig. 2, B and C). There was a close correlation between renal AOPPs levels and the number of infiltrated macrophages in glomeruli (r = 0.880; n = 72; P < 0.001) or in interstitium (r = 0.839; n = 72; P < 0.001). Intervention of apocynin significantly attenuated kidney macrophage influx induced by diabetes per se (group 5) and abolished the macrophage infiltration resulted from AOPPs challenge (group 4) (Fig. 2).
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) and TGF-β1 mRNA (Fig. 4A) was up-regulated in diabetic rats as compared with normal controls. Animals treated with AOPPs-RSA showed significantly increased MCP-1 and TGF-β1 mRNA expression when compared with RSA- or vehicle-treated diabetic rats (Figs. 3C and 4C). Protein expression of MCP-1 (Fig. 3, D and E) and TGF-β1 (Fig. 4, D and E) in renal cortex, determined by immunohistological staining, exhibited similar patterns. In control rats, MCP-1 and TGF-β1 staining was limited to minimal positive reaction of tubular cells and almost negative in glomeruli. Among diabetic animals, positive staining of MCP-1 (Fig. 3, D and E) and TGF-β1 (Fig. 4, D and E) was observed in both glomeruli and tubulointerstitium. Significantly higher staining scores were found in AOPPs- compared with RSA- or vehicle-treated diabetic rats.
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) and TGF-β1 (Fig. 4B) in diabetic rats were significantly higher than that in controls. AOPPs challenge significantly increased urinary MCP-1 and TGF-β1 levels as compared with RSA- or vehicle-treated animals. There was a close correlation between renal AOPPs levels and urinary MCP-1 (r = 0.804; n = 72; P < 0.001) or TGF-β1 excretion (r = 0.861; n = 72; P < 0.001).
Again, intervention of apocynin significantly inhibited MCP-1 and TGF-β1 expression in diabetic animals with or without AOPPs treatment. AOPPs-induced MCP-1 and TGF-β1 expression was completely prevented by apocynin (Figs. 3 and 4).
AOPPs administration increased glomerular hypertrophy, ECM expression, and albuminuria
Glomerular volume and glomerular mesangial area, determined by morphometric image analysis, were significantly increased with diabetes (Fig. 5, A and B). These parameters, reflecting glomerular hypertrophy and mesangial expansion, were significantly deteriorated in diabetic rats challenged by AOPPs-RSA as compared with those treated by vehicle or native RSA. The increase of glomerular hypertrophy was in parallel with the increase of glomerular expression of fibronectin (Fig. 5C), urinary albumin excretion, and Ccr (Table 2). Intervention of apocynin prevented AOPPs-induced glomerular hypertrophy, overexpression of fibronectin, and increase of Ccr and albuminuria.
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, O2– generation by renal cortical homogenates from diabetic rats was significantly increased compared with that from control rats. Chronic administration of AOPPs-RSA significantly enhanced O2– production by renal homogenates as compared with RSA- or vehicle-treated diabetic animals. To verify the enzymatic sources of O2– generation, production of O2– in response to NADPH was also examined in kidney homogenates in the presence of inhibitors of various enzymatic systems involved in reactive oxygen species (ROS) production. Superoxide generation was completely blocked by DPI, whereas rotenone, allopurinol, indomethacin, and L-NMMA had no effect. Intervention of apocynin significantly decreased the renal NADPH oxidase activity in diabetic rats with (group 4) or without (group 5) AOPPs challenge.
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, B and C). AOPPs administration significantly increased p47phox expression in the membrane but not in the cytosolic fraction. The ratio of p47phox expression in membrane compared with cytosol was 1.40 ± 0.03 in AOPP-, 1.24 ± 0.02 in RSA-, and 1.25 ± 0.03 in vehicle-treated diabetic animals. Apocynin intervention completely inhibited AOPPs-induced p47phox expression in the membrane and decreased the ratio of membrane p47phox and cytosolic p47phox in both AOPPs-treated (0.82 ± 0.02) and vehicle-treated (0.79 ± 0.02) diabetic rats.
The expression of membrane-bound subunit gp91phox was also enhanced in diabetic animals as compared with controls (Fig. 6D). Chronic AOPPs loading increased gp91phox expression, and this effect could be blocked by treatment with apocynin. The expression of membrane subunit p22phox was unchanged in diabetic rats and was not modified by AOPPs or apocynin (Fig. 6E).
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Discussion |
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The mechanisms that are involved in the proinflammatory effect of AOPPs remain to be clarified. In our results, plasma AOPPs levels in experimental animals increased after STZ injection, suggesting that, as reported previously, AOPPs spontaneously generated in diabetes (11, 12). Chronic administration of AOPPs in this STZ-induced diabetic model significantly increased AOPPs level in renal tissue, which was accompanied by increased production of renal ROS but not by changes of glycemia and blood pressure. Consistent with the observation, our previous study has shown that AOPPs administration induced an imbalance of redox reaction in kidneys of normal rats (14). These data suggested that AOPPs accumulation might enhance renal oxidative stress in diabetic kidney. Oxidative stress has emerged as a critical pathogenic factor in the development of diabetic nephropathy (25, 26). Although multiple pathways may result in ROS generation, recent studies indicate that a phagocyte-like NADPH oxidase is a major source of ROS in many nonphagocytic cells, including renal cells such as tubular epithelial cells and mesangial cells (27, 28, 29). Here, we provided several lines of evidence demonstrating that AOPPs promote oxidative stress and inflammatory responses in diabetic kidney mainly through activation of renal NADPH oxidase. First, AOPPs administration significantly increased NADPH oxidase-dependent O2– generation in renal homogenate as assessed by lucigenin-enhanced chemiluminescence. AOPPs-enhanced O2– production could be blocked by the NADPH oxidase inhibitor DPI (in vitro) or apocynin (in vivo), suggesting NADPH oxidase as the major source of ROS triggered by AOPPs. Second, AOPPs administration significantly enhanced renal expression of NADPH oxidase subunit p47phox in the membrane but not in the cytosolic fraction, indicating that AOPPs promoted p47phox membrane translocation, a critical step for the activation of the oxidase. The expression of another membrane-bound subunit, gp91phox, was also increased in the kidney of diabetic rats challenged by AOPPs. Moreover, enhanced renal expression of p47phox and gp91phox could be prevented by oral administration of apocynin. Apocynin is excreted in active form through the kidney (30). After cellular uptake and peroxidation, it prevents phosphorylation of p47phox, blocks p47phox association with gp91phox, and blunts NADPH oxidase activation (31, 32). Third, AOPPs-enhanced renal ROS production and tissue inflammatory responses (i.e. macrophage infiltration and overexpression of MCP-1 and TGF-β1) as well as the renal structural and functional abnormalities (i.e. glomerular hypertrophy, ECM accumulation, and albuminuria) could be prevented by intervention of apocynin, suggesting that the AOPPs-induced worsening of oxidative stress and inflammation in diabetic kidney might be associated with overactivation of renal NADPH oxidase. ROS generated during oxidative stress have been demonstrated to be signals for the activation of nuclear factor-
B, a major inflammatory transcription factor (33). The finding that AOPPs levels were closely correlated with the renal inflammatory markers (i.e. macrophage infiltration and urinary MCP-1 and TGF-β1 levels) support that AOPPs might act as a novel class of mediators for redox-sensitive inflammation in diabetic kidney. Previous in vitro study has also demonstrated that HOCl-modified low-density lipoprotein up-regulates inflammatory and fibrogenic genes in human renal tubular epithelial cells (34). Therefore, it seems reasonable to assume that in addition of hyperglycemia, chronic AOPP accumulation may constitute a new molecular basis for enhanced oxidative stress and inflammation in diabetic kidney.
In contrast to oxidized lipid, the impact of oxidized proteins in diabetes has not been extensively studied. AOPPs are prevalent in diabetes in which enhanced oxidative stress favors oxidative protein damage and therefore AOPPs formation (9, 11). Given that AOPPs accumulation occurs from an early stage of diabetes, when microangiopathy was not present (12), it is plausible to propose that AOPPs accumulation resulting from oxidative stress in diabetes can worsen the redox imbalance and inflammation. The enhanced oxidative stress and inflammation may further increase AOPPs formation via stimulation of leukocytes to produce more oxidants (34). This proinflammatory feedback loop could amplify or maintain the redox-sensitive inflammation, at least in kidney.
Conclusion
We have identified AOPPs as a new class of proinflammatory mediators in diabetic kidney. AOPPs enhance renal oxidative stress and inflammatory responses mainly through activation of renal NADPH oxidase. These data provide new information toward understanding the pathogenic basis of kidney inflammation in diabetes and may provide targets for intervention.
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Footnotes |
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Disclosure Statement: The authors have nothing to disclose.
First Published Online January 3, 2008
Abbreviations: AGEs, Advanced glycation end products; AOPPs, advanced oxidation protein products; Ccr, creatinine clearance; DPI, diphenylene iodonium; ECM, extracellular matrix; MCP-1, monocyte chemoattractant protein-1; NADPH, nicotinamide adenine dinucleotide phosphate; L-NMMA, NG-monomethyl-L-arginine; ROS, reactive oxygen species; RSA, rat serum albumin; STZ, streptozotocin.
Received November 9, 2007.
Accepted for publication December 26, 2007.
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, P < 0.001 vs. group 3.
-ED1) profiles of macrophage infiltration at wk 10. Glomerular (B) and interstitial (C) macrophage infiltration was quantified by calculation of the number of ED1-positive cells. Data are means ± SEM; n = 6 in each group at each time point. ANOVA, P < 0.001 in wk 5 and 10 in both A and B. *, P < 0.001 vs. control; #, P < 0.001 vs. groups 1 and 2; II, P < 0.01 vs. groups 1 and 2; 
