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Oxidation of cysteine and homocysteine by bovine albumin
ABB Archives of Biochemistry and Biophysics 431 (2004) 178–188 www.elsevier.com/locate/yabbi
Oxidation of cysteine and homocysteine by bovine albumin Marı´a Gabaldon* Unidad de Histoquimia, Centro de Investigacio´n, Hospital La Fe, Avenida Campanar, 21, 46009 Valencia, Spain Received 9 June 2004, and in revised form 27 August 2004
Abstract The autooxidation of cysteine and homocysteine to their disulfide forms was determined by measuring the time course of thiol groups disappearance. We found the oxidative chemistry of cysteine and homocysteine to be quite different. In the absence of added Cu(II), cysteine autooxidized at a slower rate than homocysteine, though in its presence cysteine oxidation was much faster, homocysteine being found to be a poor responder to copper catalysis. Albumin speeded up the spontaneous oxidation of both aminothiols, the reaction being faster with cysteine than with homocysteine. The copper content of different albumins was found to be highly variable, ranging from 12.75 to 0.64 lg Cu(II)/g albumin. We propose that copper bound to albumin possesses redox cycling activity to perform cysteine oxidation since: (i) copper elimination by copper chelators markedly reduces oxidation; and (ii) a positive correlation exists between the albumin copper content and the oxidation reaction rate. 2004 Elsevier Inc. All rights reserved. Keywords: Albumin; Mercaptalbumin; Cysteine oxidation; Homocysteine oxidation; Copper chelators
Albumin is a single chain multidomain protein with 17 disulfide bonds and one free thiol at Cys34, which partially forms mixed disulfides with cysteine (Cys)1 and glutathione. Albumin has a strong binding site for Cu(II) (Ka approaching 1012 M1) [1,2] at the N-terminal site involving the amino terminal nitrogen, the next two peptide nitrogens and the histidyl imidazole nitrogen forming a square planar configuration [3]. The content of Cu(II) in commercial human albumin is highly variable, which suggests that problems of metal contamination or losses arise during manufacturing processes [4]. The copper content described in a recent work was 3.87 lmol Cu(II)/mmol albumin, which indicates *
Fax: +34 6 1973018. E-mail address: [email protected]. 1 Abbreviations used: Cys, cysteine; Hcys, homocysteine; DPDS, 4,4 0 -dipyridyl disulfide; DEPC, diethyl pyrocarbonate; DTT, DL dithiothreitol; EDTA, ethylenediaminetetraacetic acid, disodium salt; DTPA, diethylenetriaminepentaacetic acid; TCA, trichloroacetic acid; Tris, Tris-(hydroxymethyl)aminomethane; BDS, bathocuproinedisulfonic acid, disodium salt; BCA, bicinchoninic acid, disodium salt; Hepes, N-(2-hydroxyethyl)piperazine-N 0 -(2-ethanesulfonic acid). 0003-9861/$ - see front matter 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2004.08.028
that only 0.4% of the albumin molecules carry bound copper [5]. The present study was stimulated by the observation that when preparing aged albumin the –SH groups of Cys and homocysteine (Hcys) completely disappeared after 24 h of incubation with albumin at pH 8.9 and low ionic strength. The fact that the complete oxidation of Cys also occurred when increasing the Cys/albumin molar ratio from 5 to 20 pointed to a catalytic phenomenon where a metal with redox cycling activity, most probably copper, catalyzed Cys oxidation. This observation led us to hypothesize that, under physiological conditions, in the relatively oxidative environment of blood, albumin could play a critical role in the oxidation of Cys and Hcys to their disulfide forms through the catalytic action of the minimal amounts of Cu(II) that it binds and transports. In fact, aminothiols circulate mainly as oxidized forms and only 0.5% of total Hcys and 5% of total Cys are found in the reduced form [6]. Aminothiols in human plasma exist in reduced and free and albumin-bound oxidized forms interacting with each other through redox and disulfide exchange
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reactions. The aminothiols together with the free –SH group of albumin comprise a dynamic system referred to as the redox thiol status and represent the balance between prooxidant and antioxidant processes in plasma [7]. This balance may modulate redox-sensitive signaling pathways and suffers a marked prooxidative shift with aging and in cardiovascular and degenerative diseases [7,8]. The regulation of these redox systems in plasma has been little studied. Recently, Sengupta et al. [5,9] proposed that in circulation, albumin mediates the oxidation of Hcys by thiol disulfide exchange, whereas ceruloplasmin drives Cys oxidation by copper-dependent oxidative mechanisms. The basic question posed in this study is whether the copper bound to albumin has redox-cycling activity to perform the oxidation of aminothiols. We first studied the autooxidation of Cys and Hcys in the absence of albumin, and the response of this reaction to copper catalysis. We found that despite the structural similarity, the chemical behavior was quite different, the oxidation of Hcys being much faster and much less sensitive to copper catalysis than that of Cys. To know whether the effect we first observed was a general phenomenon we studied albumins with different compositions and purities to exclude the possible effect of copper-rich contaminant globulins. All studied albumins showed a prooxidant effect on Cys and this effect was related to the copper content of the albumin. Finally, in order to demonstrate that copper bound to albumin was involved in Cys oxidation, experiments were performed after removing copper with different types of copper chelators. Results indicate that albumin may play a crucial role in Cys oxidation by a copper-catalyzed oxidative process and that copper-dependent Hcys oxidation by albumin is a minor pathway in the overall conversion to disulfide forms. As proposed by Sengupta et al. [5], a thiol disulfide exchange with the disulfide bond at Cys34 of albumin seems to be a more plausible mechanism for Hcys oxidation.
Materials and methods Chemicals The following bovine serum albumins were supplied by Sigma Chemical:A-4503 lot 050K0891 remaining most globulins; A-0281 essentially fatty acid and globulin free; A-7638 essentially globulin free; A-7511 essentially fatty acid free; and A-3912 remaining most globulins and obtained by heat shock. The following bovine serum albumins were supplied by ICN Biomedicals: 194774 essentially globulin free and 810012 crystalline. Bicinchoninic acid disodium salt, bathocuproinedisulfonic acid disodium salt, L -cysteine free base (C-7352), DL -homocysteine, L -histidine (H-8000), diethylenetri-
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aminepentaacetic acid (D-6518), DL -dithiothreitol (D0632), iodoacetamide (I-6125), L -ascorbic acid (A-5960), diethyl pyrocarbonate, and trichloroacetic acid (T9159) were supplied by Sigma Chemical Co. Copper atomic spectroscopy standard solution and 4, 4 0 -dipyridyl disulfide (DPDS; Aldrithiol-4) were supplied by Fluka and Aldrich, respectively. Solutions were prepared in deionized water obtained with an Elix water purification system from Millipore (10–15 MX cm resistivity). Buffered albumin solutions were prepared as previously described [10]. Determination of thiol groups Thiol groups were determined with DPDS as described by Riener et al. [11] with some modifications. The DPDS reagent is a 4 mM DPDS solution in 12 mM HCl and was prepared by dissolving 45 mg of DPDS in 1.5 ml of 0.4 M HCl before completing with H2O to 50 ml. Aliquots were immediately frozen at 20 C and thawed portions were used within 1 h. Reaction was performed by mixing 1.2 ml of the study sample in 0.1 M sodium phosphate buffer pH 6 with 50 ll of the DPDS reagent. Albumin concentration for thiol group determination must be between 0.5 and 5 mg/ml and aminothiol concentration between 4 and 40 lM. A reagent blank with buffer instead of study sample and a sample blank with buffer instead of DPDS were processed in the same way. Absorbances at 324 nm were measured against H2O after 10 min. The thiol group reacts with DPDS to produce an equivalent of 4-thiopyridone, which has a molar absorptivity of 21,400 M1 cm1 at 324 nm. The concentration of 4-thiopyridone was calculated from the absorbance at 324 nm after subtracting the absorbances of the two blanks. Determinations were performed in duplicate. Size exclusion chromatography In some experiments albumin was incubated with low molecular weight reagents, which after reaction must be completely removed by gel filtration. The separation of albumin from low molecular weight products was performed by passing solutions through a Hi Prep 26/10 desalting column (Amersham Biosciences) equilibrated with H2O. A 10-ml sample of A-4503 albumin filtered through 0.45 lm was applied at 5 ml/min followed by 1 ml washing with H2O; elution was performed with H2O at 2.5 ml/min and the albumin fraction was collected between 2 and 8 min after the start of elution. Given that the molar concentration of the reagents considerably exceeded that of albumin, controls were performed to make sure that the eluate fraction with albumin was free from low molecular weight products. The pattern of histidine elution was obtained from a 45-mM solution by monitoring the conductivity in
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eluates or by histidine determination with diethyl pyrocarbonate (DEPC). In this latter case 600 ll of eluate were mixed with 580 ll of 0.1 M sodium phosphate buffer pH 7.5 and with 20 ll of 91 mM DEPC in acetonitrile prepared as indicated in [10]. Readings were taken 15 min later at 240 nm. The pattern of DL -dithiothreitol (DTT) elution was obtained by determination of the thiol groups in eluates after applying a 1.8 mM DTT solution. Iodoacetamide, ethylenediaminetetraacetic acid (EDTA), and diethylenetriaminepentaacetic acid (DTPA) could be determined by monitoring the conductivity of the effluents. All the determinations showed the fraction collected between 2 and 8 min under the described conditions to contain albumin without the presence of low molecular weight substances.
thione. A 72 mg/ml A-4503 albumin solution in 0.3 M NaCl–0.1 M sodium phosphate buffer (final pH 6.85) (10 ml) was mixed with a 10.8 mM DTT solution in 0.3 M NaCl–0.1 M sodium phosphate buffer pH 6.75 (2 ml), keeping the final pH between 6.76 and 6.84. The reaction mixture was filtered through a 0.45-lm filter, and after 25 min of incubation a 10 ml sample was applied to a Hi Prep 26/10 desalting column under the conditions previously described. The collected albumin fraction was analyzed for thiol groups after diluting 1/ 20 with 0.1 M sodium phosphate buffer pH 6. The – SH/albumin molar ratio was found to be 1.00 ± 0.05 (mean ± SD, n = 4). The composition of mercaptalbumin determined by gel filtration chromatography was the same as the starting albumin (86.1% monomer, 11.8% dimer, and 2.1% polymers).
Autooxidation of aminothiols Blocking of albumin –SH groups The reaction mixture at pH 7.4 contained: aminothiol, 1.5 mM; Tris–HCl buffer pH 7.5, 10 mM; and NaCl, 150 mM. The reaction mixture at pH 8.9 was: aminothiol, 1.5 mM; and Tris–HCl buffer pH 9, 50 mM. The 25 mM starting aminothiol solutions were freshly prepared. Incubations were performed for 24 h at room temperature in the dark. Samples were taken at 2, 4, 8, and 24 h, diluted (1/40 to 1/10) with 0.1 M sodium phosphate buffer pH 6, and thiol groups were determined by DPDS analysis. Values at t = 0 were obtained by diluting the starting aminothiol solution with H2O to 1.5 mM. After 1/40 dilution with 0.1 M sodium phosphate buffer pH 6 and 4 min of incubation, samples were taken for thiol determination. Oxidation of aminothiols by albumin The reaction mixture contained: albumin, 300 lM; NaCl, 150 mM; and aminothiol, 1.5 mM; pH adjusted to 7.4 with 0.2 M Tris. Aminothiol solutions were freshly prepared. Albumin solution was filtered through a 0.45-lm filter. Incubations, values at t = 0, and DPDS analysis were performed as indicated in the previous section. Total thiol groups in the reaction mixture were the sum of the added thiol groups (1500 lM) and the thiol groups carried by the albumin. The –SH/albumin molar ratio of albumins ranged from 0.40 to 0.55, and was calculated by determination of the thiol groups in a 60 lM albumin solution in 0.1 M sodium phosphate buffer pH 6. The concentration (lM) of the albumin thiol groups was obtained by multiplying 300 by the –SH/albumin molar ratio. Preparation of mercaptalbumin Mercaptalbumin has the thiol group at Cys34 fully reduced whereas normal albumin has close to 50% of this thiol group forming disulfide bonds with Cys and gluta-
The free –SH group of albumin was blocked with iodoacetamide. The final concentrations in the reaction mixture were: albumin, 900 lM; iodoacetamide, 4.5 mM; NaCl, 50 mM; final pH adjusted with 0.2 N NaOH, 7.5. Incubation was carried out for 1 h in the dark, and the solution was passed through a Hi Prep 26/10 desalting column under the conditions previously described. The collected albumin fraction was negative for thiol groups. Copper calibration curve A 1 mM Cu(NO3)2–6 mM glycine stock solution was prepared from a copper atomic spectroscopy standard solution. Copper–glycine stock solutions were used to prevent the formation of metal-hydroxy and metal-oxy polymers in neutral media [12]. Copper(II) was determined after reduction by ascorbic acid by spectrophotometric analysis at 358 nm of the Cu(I) complex formed with bicinchoninic acid (BCA). As BCA from different suppliers showed important differences in capacity to complex Cu(I), before starting determinations BCA must be assayed by adding increasing amounts to a fixed copper concentration covering a BCA/Cu(I) molar ratio range from 2 to 3 at 0.2 intervals, until absorbances are near constant. As the molar absorptivity of the BCA– Cu(I) complex is approximately 23 times greater than that of BCA, absorbance increases notably when increasing BCA concentration if this is defective, and very discretely when in excess. In our case the BCA/ Cu(I) molar ratio for complete copper complexation was 2.3. Calibration curves were prepared under the conditions described for the analysis of copper in albumin (section below), except that BCA solution in 0.6 M sodium Hepes–0.15 M aOH was used instead of 0.6 M Hepes buffer pH 8. In all cases the final pH was 6.8–7.2, and the BCA/Cu(I) molar ratio was 2.3. Extinc-
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tion coefficients were calculated from calibration curves after subtracting absorbances of blanks without copper and BCA from those of the study samples. Under these conditions, the extinction coefficient of the BCA–Cu(I) complex at 358 nm was 45.37 ± 1.62 mM1 cm1 (mean ± SD, n = 23), and that of BCA was found to be 1.94 ± 0.28 mM1 cm1 (mean ± SD, n = 23). Copper determination in albumin The copper bound to albumin was determined by a modification of the technique described by Brenner and Harris [13], based on the separation of copper(II) in strong acid media, reduction to Cu(I) with ascorbic acid, and formation of a complex with BCA, which is then spectrophotometrically assayed. An albumin solution was prepared by mixing 1.2 g of albumin and 3.6 ml of H2O in a 10-ml vial with magnetic stirring. After 1/160 dilution, readings at 280 nm were taken and albumin concentration was adjusted to 3.5 mM (extinction coefficient of albumin at 280 nm, 65,500 M1 cm1). Albumin solution was mixed with 30% trichloroacetic acid (TCA) in a 3/1 ratio v/v. The suspension was stirred in a 10-ml vial for 2 h and centrifuged two times at 5000 rpm for 20 min. The supernatant obtained under these conditions represented 60% of the initial suspension volume. Copper determination was performed by mixing in the order stated: 0.60 ml of supernatant, 0.12 ml of freshly prepared 2 mM ascorbic acid, and 0.48 ml of 60 or 100 lM BCA in 0.6 M Hepes buffer pH 8. A blank without BCA was processed in the same way. Readings were taken at 358 nm against H2O, and the absorbance of the blank was subtracted from that of the study sample. As BCA forms a 2:1 complex with Cu(I), BCA concentration must be greater than double that of Cu(I) but not too high to minimize interferences produced by excess BCA, which also absorbs, though discretely, at 358 nm. To conciliate these two requirements, BCA concentration has been adapted to that of Cu(I) in the following way: beginning with a 60 lM BCA concentration, absorbance of the study sample minus the blank must be <0.5; if absorbance >0.5, then a 100 lM BCA concentration must be used. When absorbance is >0.88 with this last concentration, the supernatant must be diluted 1/2 with 4.5% TCA. This scheme covers a range of copper concentrations in the supernatant from 6 to 80 lM. The absorbance of the reaction mixture is the sum of the absorbances of the BCA–Cu(I) complex and excess BCA. Copper concentration in the reaction mixture (lM) was calculated as =(a b * c)/(d 2b) where a = absorbance of the study sample minus that of the blank; b = lM absorptivity of BCA (1.94 · 103); c = initial BCA concentration in the reaction mixture (lM); and d = lM absorptivity of the BCA–Cu(I) complex (45.37 · 103). Copper concentration in the starting 3.5 mM albumin
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solution was calculated by multiplying copper concentration in the reaction mixture by 2 and dividing by 0.75/0.6. Determinations were performed in duplicate. Copper reduction assay in the absence of albumin Aminothiols in the presence of added Cu(II) oxidize and produce Cu(I). Hcys oxidation by copper was studied at different molar ratios by monitoring the formation of Cu(I) with bathocuproinedisulfonic acid (BDS) or BCA. The reaction mixture contained: Cu(II) (40 lM Cu(NO3)2–240 lM glycine), 40 lM; sodium phosphate buffer pH 7.4, 20 mM; NaCl, 150 mM; BDS, 120 lM; and freshly prepared Hcys, 400 or 40 lM for a Hcys/ Cu(II) molar ratio of 10 or 1, respectively. When using BCA, final concentrations of BCA and Hepes buffer pH 7.4 were 92 lM and 20 mM, respectively. Reaction was started by addition of Hcys. Copper reduction was monitored by registering the absorbance of the complex BDS–Cu(I) at 480 nm, considering an absorptivity of 12,540 M1 cm1 [14]. When using BCA, Cu(I) in the reaction mixture was determined by absorbance at 358 nm after applying the formula described in the previous section. To correlate formation of Cu(I) and disappearance of thiol groups at different aminothiol/Cu(II) molar ratios, incubation mixtures were prepared containing: Cu(II) (40 lM Cu (NO3)2–240 lM glycine), 40 lM; sodium phosphate buffer pH 7.4, 20 mM; NaCl, 150 mM; and freshly prepared aminothiol, 400 or 40 lM for a aminothiol/Cu(II) molar ratio of 10 or 1, respectively. Reaction was started by addition of the aminothiol, and after 5 min samples were taken for DPDS analysis. When using an aminothiol/Cu(II) molar ratio of 10, samples were diluted 1/10 with 0.1 M sodium phosphate buffer pH 7.4, while the samples were used undiluted when the molar ratio was 1. Values at t = 0 were obtained by diluting the starting aminothiol solution with 20 mM sodium phosphate buffer pH 7.4 to 40 lM. Copper reduction assay in the presence of albumin Since techniques for Cu(I) determination are not sufficiently sensitive to detect the low amount of copper bound to albumin, a copper enriched albumin with 50% occupation of its Cu(II) binding site was prepared by the addition of exogenous Cu(II). Binding of Cu(II) to albumin was performed by incubating for 50 min equal volumes of 150 lM Cu(II) (150 lM Cu(NO3)2 and 900 lM glycine) and 300 lM of A-4503 albumin in 0.1 M sodium phosphate buffer, final pH 7.4. For assessing complete copper binding to albumin, an aliquot was filtered through a 30 K Omega-type Filtron membrane in an Amicon 50 ml ultrafiltration cell, and the Cu(II) content was analyzed in the filtrate. Similar incubation with buffer instead of albumin was carried
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out. The reaction mixture contained the following: NaCl, 150 mM; albumin, 100 lM; Cu(II) (50 lM Cu(NO3)2–300 lM glycine), 50 lM; BDS, 150 lM; and freshly prepared aminothiol, 500 lM. Reaction was started by addition of aminothiol. Copper reduction was monitored by registering the absorbance of the complex BDS–Cu(I) at 480 nm, considering an absorptivity of 12,540 M1 cm1. Similar reaction mixtures were prepared with the filtrate obtained by passing the Cu(II) enriched albumin through a 30 K membrane. The net absorbances were calculated by deducting the absorbances of the blanks prepared without thiol from the absorbances of the study samples. Disulfide reduction assay To demonstrate that aminothiols in the presence of albumin were oxidized to their disulfide forms, disulfide bonds were reduced by DTT, and the formed thiol groups were determined after complexation of excess DTT by arsenite [15]. Albumin (300 lM) was incubated with Cys or Hcys (1500 lM) in 150 mM NaCl, pH 7.4 for 24 h and was then filtered through a Filtron membrane 30 K Omega-type in an Amicon 50 ml ultrafiltration cell. Filtrate was diluted 1/2, adjusting the final pH to 8 with Tris–HCl buffer pH 8. This solution was mixed with an equal volume of 3.8 mM DTT–1 mM EDTA in 50 mM Tris–HCl buffer pH 8, and incubated for 70 min under nitrogen. Reaction mixture was diluted 1/10 with 3.8 mM sodium arsenite in 0.1 M sodium phosphate buffer pH 6 and incubated for 8 min. Thiol groups were immediately determined by DPDS reaction. A blank with 20 mM Tris–HCl buffer pH 8 instead of filtrate was processed in the same way, and its absorbance subtracted from that of the study samples. Analytical procedures Fractionation of albumin in monomeric and polymeric forms was performed by gel filtration using Superdex 200 prep grade on a HiLoad 16/60 column (Amersham Biosciences). The column was loaded with 1 ml 20 mg/ml albumin solution and eluted with 0.15 M NaCl in 10 mM sodium phosphate buffer pH 7.4 at a pumped flow rate of 0.8 ml/min.
Results Autooxidation of aminothiols Fig. 1A shows the effect of pH on the time course of aminothiol oxidation measured by the loss of –SH groups in mixtures without added copper. Under physiological conditions (pH 7.4 and 150 mM NaCl) the rate of autooxidation is faster for Hcys than for Cys. This
Fig. 1. Autooxidation of aminothiols. Time course of –SH group disappearance. (A) Effect of pH. The reaction mixture at pH 7.4 contained: aminothiol, 1.5 mM; Tris–HCl buffer pH 7.5, 10 mM; and NaCl, 150 mM. The reaction mixture at pH 8.9 was: aminothiol, 1.5 mM; and Tris–HCl buffer pH 9, 50 mM; Cys pH 7.4 (——); Cys pH 8.9 (—j—); Hcys pH 7.4 (—m—); Hcys pH 8.9 (—d—). (B) Effect of Cu(II). The reaction mixture contained: aminothiol, 1.5 mM; Tris–HCl buffer pH 7.5, 10 mM; NaCl, 150 mM; and Cu(II) (0.61 lM Cu(NO3)2–3.67 lM glycine), 0.61 lM. Final pH, 7.4; Cys (——); Cys + Cu (II) (—d—); Hcys (—m—); Hcys + Cu(II) (—j—). The amount of remaining –SH groups was determined by DPDS analysis, as described in Materials and methods. Each point represents the mean from five independent experiments. The coefficient of variation did not exceed 20%.
may be explained taking into account that Hcys is a stronger nucleophile than Cys, and is thus more reactive [16]. The rise in pH to 8.9 practically does not modify the rate of Cys oxidation but drastically increases that of Hcys due to the lower pKa of the thiol group of the former [9]. Fig. 1B shows the time course of the oxidation of aminothiols in the presence of added Cu(II) measured by the loss of –SH groups. A low copper concentration (0.61 lM) has been used in order to compare the effect of free Cu(II) added to the reaction mixture with the effect of Cu(II) bound to commercial albumin described in the following section in which the same final copper concentration was obtained from a 300 lM A-4503 albumin solution (Table 1). Copper(II) was much more efficient in catalyzing Cys oxidation than Hcys oxidation. At 2 h, Cys lost 1% of its –SH
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Table 1 Cysteine oxidation and albumin copper content Albumin
Loss of –SH groups (%)a
Copper contentb lgCu(II)/g albumin
2h
4h
8h
24 h
A-4503 810012 A-3912
46.6 94.9 83.3
74.8 97.9 95.9
93.9 99.2 98.6
98.4 99.7 99.5
1.95 12.75 2.33
Globulin free A-7638 194774
40.1 89.4
64.1 96.4
88.6 98.5
98.0 99.6
1.44 4.42
Fatty acid free A-7511 A-0281
38.9 15.6
62.3 31.6
86.8 52.4
96.7 90.8
0.73 0.64
a The reaction mixture contained: albumin, 300 lM; NaCl, 150 mM; and Cys, 1.5 mM; pH adjusted to 7.4 with 0.2 M Tris. The amount of the remaining –SH groups was determined by DPDS analysis, as described in the ‘‘Materials and methods’’ section. Each value represents the mean from three independent experiments. The coefficient of variation did not exceed 15%. b Each value represents the mean from three independent determinations. The coefficient of variation did not exceed 10%.
groups in the absence and 94% in the presence of Cu(II), whereas the losses in –SH groups for Hcys at the same time were 7 and 17%, respectively. Oxidation of aminothiols in the presence of Cu(II) can be determined by measuring the disappearance of thiol groups or the formation of Cu(I) with chelating reagents such as BCA or BDS. The time course of Hcys oxidation was different depending on the reagent used for determination of the formed Cu(I) (Fig. 2). At a Hcys/Cu(II) molar ratio of 10, the kinetics with BCA was characterized by a lag-phase of 15 min followed by a period where Cu(I) was formed at a slow rate. The reaction was dependent on the buffer used, being
Fig. 2. Autooxidation of Hcys by Cu(II). Time course of Cu(I) formation. The reaction mixture contained: Cu(II) (40 lM Cu (NO3)2– 240 lM glycine), 40 lM; NaCl, 150 mM; BCA or BDS, 92 lM or 120 lM, respectively; Hepes buffer or sodium phosphate buffer pH 7.4, 20 mM; and Hcys, 40 or 400 lM. Reaction was started by addition of Hcys. Numbers in parentheses indicate the Hcys/Cu(II) molar ratio. BCA (1) (—m—); BCA (10) (- -m- -); BCA (10) with phosphate buffer (—d—); BDS (1) (——); BDS (10) (—j—). The amount of formed Cu(I) was determined by absorbance at 480 and 358 nm when using BDS and BCA, respectively, as described in Materials and methods. Each point represents the mean from three independent experiments.
faster with phosphate than with Hepes buffer at the same pH of 7.4. When using BDS, the kinetics was completely different, the reduction of Cu(II) being almost instantaneous. Other studies described a marked stimulation of copper-induced oxidation of LDL by BDS [17]. A possible explanation for these observations is that BDS also binds Cu(II), and this complex is a stronger oxidant than Cu(II) [14,18]. Thus, BDS must be used with caution to monitor Cu(II) reduction, because thiol oxidation measured in its presence could generate methodological artifacts. When Cys was incubated with Cu(II) at molar ratios of 1 and 10, the reduction of Cu(II) was almost instantaneous regardless of whether BDS or BCA was used for determination of the formed Cu(I) (data not shown). An unexpected finding of this study was the influence of the Hcys/Cu (II) molar ratio on the time course of the reaction when using BCA. At a Hcys/Cu(II) molar ratio of 1, copper reduction was almost instantaneous, whereas the reaction rate was much slower at a Hcys/ Cu (II) of 10 (Fig. 2). The same results were obtained when Hcys oxidation was measured by disappearance of thiol groups. Fig. 3 shows the oxidation of aminothiols after 5 min of incubation with Cu(II), expressed as the disappearance of thiol groups or the appearance of Cu (I). Results indicate that when using BCA for determination of Cu(I), both criteria were concordant except for a minor imbalance found at Hcys/Cu(II) molar ratio of 10 that we are unable to account for. It has recently been reported [19] that the interaction between Cu(II) and Hcys at molar ratio 1:1 leads to the formation of a blue complex with Cu (II), and that in the presence of excess ligand the complex is yellow with copper in the reduced state. The differences we have observed at Hcys/Cu(II) molar ratios of 1 and 10 may be related to the oxidative capacity of the different Hcys/copper complexes formed.
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Fig. 3. Autooxidation of aminothiols by Cu(II). –SH group disappearance and Cu(I) formation at 5 min For determination of the remaining –SH groups, the reaction mixture contained: Cu(II) (40 lM Cu(NO3)2–240 lM glycine), 40 lM; NaCl, 150 mM; sodium phosphate buffer pH 7.4, 20 mM; and aminothiol, 40 or 400 lM. Reaction was started by the addition of aminothiol. For determination of the Cu(I) formed, the reaction mixture contained: Cu(II) (40 lM Cu(NO3)2– 240 lM glycine), 40 lM; NaCl, 150 mM; BCA, 92 lM; Hepes buffer, pH 7.4, 20 mM; and aminothiol, 40 or 400 lM. Reaction was started by the addition of aminothiol. Numbers indicate the aminothiol/Cu(II) molar ratio. Dotted bars, disappearance of –SH groups. Dashed bars, formation of Cu (I). The amount of formed Cu(I) was determined with BCA by measuring the absorbance at 358 nm, and the amount of remaining –SH groups was determined by DPDS analysis, as described in Materials and methods. Values plotted are the mean from three independent experiments.
Oxidation of aminothiols by albumin Fig. 4 shows the time course of the oxidation of aminothiols by albumin at pH 7.4, measured by the loss of thiol groups. Albumin induces a loss of –SH groups in both aminothiols, the reaction rate being faster for
Fig. 4. Oxidation of aminothiols by albumin. Time course of –SH group disappearance. The reaction mixture contained: albumin (A-4503), 300 lM; NaCl, 150 mM; and aminothiol, 1.5 mM; pH adjusted to 7.4 with 0.2 M Tris; Cys (——); Cys + albumin (—d—); Hcys (—m—); Hcys + albumin (—j—). The amount of the remaining –SH groups was determined by DPDS analysis, as described in Materials and methods. Each point represents the mean from five independent experiments. The coefficient of variation did not exceed 20%.
Fig. 5. Aminothiol oxidation by albumin. Time course of Cu(I) formation. The reaction mixture contained: albumin (A-4503), 100 lM; Cu(II) (50 lM Cu (NO3)2–300 lM glycine), 50 lM; NaCl, 150 mM; aminothiol, 500 lM; BDS, 150 lM; sodium phosphate buffer, pH 7.4, 20 mM; albumin and Cu(II) were incubated for 50 min before addition of the remaining reagents. Reaction was started by the addition of aminothiol; Cys (—j—); Cys + albumin (——); Hcys (—m—); Hcys + albumin (—d—). The amount of Cu(I) formed was determined by absorbance at 480 nm, as described in Materials and methods. Each point represents the mean from three independent experiments.
Cys than for Hcys. Thiol groups disappearance was due to the formation of disulfide bonds since 95–100% of the initial –SH groups were recovered from the filtrate through a 30 K membrane after DTT reduction and complexing excess DTT by arsenite. To examine whether the copper bound to albumin had redox cycling activity, aminothiols were incubated with a Cu(II) enriched albumin (albumin/Cu(II) molar ratio of 2), and the formed Cu(I) was assayed with BDS. Under the conditions used complete copper binding to albumin was produced, since the filtrate from a 30 K membrane was negative for copper; results thus refer to the effect of copper bound to albumin and not to any residual free copper. Fig. 5 shows the time course of the formation of Cu(I). In the presence of free Cu(II) and in the absence of albumin, the reduction of Cu(II) by aminothiols was nearly instantaneous. In the presence of albumin, the reduction of Cu(II) was delayed with both aminothiols, but was much more faster with Cys than with Hcys. Results indicate that copper bound to albumin has redox cycling activity, and that aminothiols reduce free Cu(II) faster than Cu(II) bound to albumin. To study the role of the albumin thiol group at Cys34 in mediating the oxidation of Cys, we performed incubations with mercaptalbumin and with albumin with the free –SH group blocked with iodoacetamide. As shown in Fig. 6, the time course of the reaction was similar in the presence and in the absence of free thiol groups, which indicates that the thiol group of albumin is not involved in the oxidation of Cys. As albumin A-4503 is a fraction V remaining most globulins there was the possibility that contaminant pro-
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EDTA. When the EDTA–Cu(II) complex was not removed from the incubation mixture, inhibition was not produced, and Cys oxidation at 2 h was 47% greater than that seen with albumin, possibly due to the greater accessibility of Cys to the copper bound to EDTA than to the copper bound to albumin (Fig. 7A). We tried to remove copper from albumin by incubating the latter with a large excess of histidine, since histidine forms chelates with Cu(II). When before beginning reaction albumin was incubated with histidine in a 1/50
Fig. 6. Oxidation of Cys by albumin. Effect of the albumin thiol group. Time course of –SH group disappearance. The reaction mixture contained: albumin A-4503, mercaptalbumin or –SH blocked albumin, 300 lM; NaCl, 150 mM; and Cys, 1.5 mM; pH adjusted to 7.4 with 0.2 M Tris; Cys (——); Cys + albumin (—j—); Cys + mercaptalbumin (—m—); Cys + albumin (–SH blocked) (—d—). The amount of the remaining –SH groups was determined by DPDS analysis, as described in Materials and methods. Each point represents the mean from five independent experiments. The coefficient of variation did not exceed 15%.
teins, mainly ceruloplasmin, played a significant role in the oxidation. Additional studies carried out with globulin-free albumins and with the purest albumins showed that Cys oxidation was produced in the absence of globulins, and that the reaction rate was variable and independent of albumin purity (Table 1). The molecular weight of ceruloplasmin (132 kDa) is similar to that of albumin dimer (133 kDa). Gel filtration completely resolved the monomeric and dimeric forms of albumin, albumin monomer can thus be considered to be free from any contaminant ceruloplasmin. Albumin A-4503 monomer oxidized Cys at a higher rate than unfractionated albumin, the disappearance of thiol groups being 94 and 47% at 2 h for monomer and unfractionated albumin, respectively. This high reactivity indicated that Cys oxidation was due to albumin itself and not to contaminant high molecular weight substances. Oxidation of cysteine by albumin: Effect of chelating agents To determine whether the copper bound to albumin played a role in Cys oxidation, assays were performed in the presence of chelating agents. Incubations in the presence of 1 mM EDTA failed to inhibit Cys oxidation (data not shown). However, when before beginning the reaction albumin was incubated with EDTA in a 1/10 molar ratio at pH 7.4 for 20 h and the EDTA–Cu(II) chelate was removed by gel filtration, the reaction rate was considerably slowed and Cys oxidation was inhibited by 91 and 43% at 2 and 24 h, respectively The same results were obtained when using DTPA instead of
Fig. 7. Oxidation of Cys by albumin. Time course of –SH group disappearance. (A) Effect of chelating agents. The reaction mixture contained: albumin (A-4503), 300 lM; NaCl, 150 mM; and Cys, 1.5 mM; pH adjusted to 7.4 with 0.1 M Tris. Albumin was preincubated with EDTA or DTPA in molar ratio 1/10 for 20 h at pH 7.4 and low molecular weight components removed by gel filtration; Cys (——); Cys + albumin (- -j- -); Cys + albumin incubated with EDTA (—j—); Cys + albumin incubated with EDTA (without desalting) (—d—); Cys + albumin incubated with DTPA (—m—). (B) Other effectors. The reaction mixture contained of albumin (A-4503), 300 lM; NaCl, 150 mM; and Cys, 1.5 mM; pH adjusted to 7.4 with 0.1 M Tris. BDS (100 lM) was added after Cys. Albumin was preincubated with histidine in molar ratio 1/50 at pH 7 for 20 h and low molecular weight components removed by gel filtration. Albumin was preincubated with HNO3 at pH 1.8 for 20 h and passed through a desalting column; Cys (——); Cys + albumin (- -j- -); Cys + albumin incubated with histidine (—m—); –cys + albumin incubated with HNO3 (—j—); Cys + albumin + BDS (—d—). The amount of remaining –SH groups was determined by DPDS analysis as described in Materials and methods. Each point represents the mean from five independent experiments. The coefficient of variation did not exceed 20%.
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molar ratio at pH 7 for 20 h and the histidine–Cu(II) complex was removed by gel filtration, the reaction rate was considerably slower than that of untreated albumin, and Cys oxidation was inhibited by 96 and 75% at 2 and 24 h, respectively (Fig. 7B). An increase in the histidine/ albumin molar ratio to 90 did not modify the results. However, when incubation with histidine was performed at pH 5, the rate of Cys oxidation was increased, reaching intermediate values between the untreated and treated albumin at pH 7 (data not shown). This pH dependency may be explained by the lower copper binding capacity of histidine at pH 5 than at pH 7 [20]. For removing copper from albumin, apart from these metal binding strategies with low molecular weight chelators, we used acid conditions, since binding of copper to albumin increases linearly from pH 4 to 7, and is minimal at lower pH values [20]. When before beginning the reaction albumin was incubated with HNO3 at pH 1.8 for 20 h and passed through a desalting column, the reaction rate was considerably slower than that of untreated albumin and Cys oxidation was inhibited by 91 and 71% at 2 and 24 h, respectively (Fig. 7B). As Cys oxidation was coupled to a continuous copper redox cycling process, we tried to inhibit reaction by avoiding reoxidation of the formed Cu(I) by atmospheric oxygen. BDS was added after Cys to the reaction mixture in order to form a redox inactive complex with Cu(I). Cys oxidation was inhibited by 68 and 44% at 2 and 24 h, respectively. Results indicate that the oxidation of Cys is coupled to the reoxidation of Cu(I) in albumin, as the reaction rate decreases when the availability of Cu(I) for redox cycling is impaired by BDS complexing (Fig. 7B). Albumin copper content As is shown in Table 1, the copper content of different commercial albumins (fraction V, crystalline, fatty acid free, and globulin free) was highly variable, and was not correlated to the presence of globulins. Elimination of globulins was not followed by a decrease in copper content, which indicates that copper is afforded by albumin itself and not by contaminant copper-rich globulins. The purest albumin (810012) had the greatest copper content (12.75 lg Cu(II)/g albumin), representing 1.3% occupancy of the copper-binding site. Contrarily, fatty acid free albumins (A-7511 and A-0281) were the poorest, with 0.73 and 0.64 lg Cu(II)/g albumin, which represents 0.08 and 0.07% of albumin molecules with bound copper, respectively. As indicated by the manufacturer, fatty acid free albumins were obtained by the method of Chen [21], which is based on the removal of fatty acids by adsorption on charcoal in strong acid medium. This condition does not favor copper binding to albumin, and apart from fatty acids the treatment can remove some of the bound copper.
Table 1 shows the time course of thiol group disappearance after incubation of different commercial albumins with Cys. At 2 h there was a positive correlation between albumin copper content and percentage thiol group loss (n = 6, r = 0.86, P < 0.05). Albumin 810012 has been excluded from this calculation, because its reaction rate was much higher than that of the rest of albumins, having lost 80% of the thiol groups at 40 min, and almost 100% at 2 h.
Discussion The oxidation of aminothiols in the presence of copper is a complex phenomenon due to interaction between copper and thiols with the formation of chelates. In the case of Cys it has been suggested that the reaction catalyst is not the free metal but a 1:2 copper–thiol complex, where the close location of the free amino group and thiol group is crucial for thiol oxidation [22]. According to this, formation of the aminothiol–copper complex would be favored in the case of Cys, due to its shorter chain length and this would explain why Hcys despite oxidizing at a greater rate than Cys in the absence of copper, would oxidize at a slower rate in its presence. Our results help explain the apparently contradictory observations of previous reports on the effect of Cu(II) upon aminothiol oxidation. When using a high aminothiol/Cu(II) molar ratio and measuring oxidation by disappearance of the thiol groups, some authors reported Cys oxidation to be much faster than Hcys oxidation, [5,23], whereas the opposite was found by other investigators when aminothiol/Cu(II) was <1 and oxidation was measured by Cu(I) assay [12,24]. In relation to these results, it has been shown that the lag times for oxidation of low density lipoprotein lipids in the presence of Cu(II) vary markedly as a function of Hcys concentration: a Hcys/Cu(II) molar ratio <1 shortens the lag time, while a molar ratio > 1 slows oxidation [25]. All these results indicate that Hcys oxidation in the presence of copper is dependent on their relative concentrations, and this fact must be taken into account when extrapolating data obtained by experimental protocols to the in vivo situation, where the actual molar ratio of the components may be quite different. We found the time course of Cys oxidation to be similar in the presence and in the absence of free –SH groups, which indicates that the albumin thiol group is not involved in Cys oxidation. These results are in discordance with those obtained by Sengupta et al. [5], who reported complete inhibition of the oxidation of both aminothiols when using mercaptalbumin. The procedure followed by these authors to obtain mercaptalbumin (DTT/albumin molar ratio of 5, 45 min incubation, and prolonged elimination of DTT by dialysis) favored
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copper removal from albumin more than our conditions (DTT/albumin molar ratio of 2, 25 min incubation, and rapid DTT elimination by desalting), and, in fact, the mercaptalbumin obtained by these authors had 94% less copper than the starting albumin. The purpose of this study was to demonstrate whether the minimal amount of copper bound to albumin under physiological conditions would be able to catalyze the oxidation of aminothiols by a redox cycling mechanism. Results indicate that copper bound to albumin exhibits a remarkable catalytic redox-cycling activity towards Cys since: (i) elimination of Cu(II) by copper chelators markedly decreases oxidation; (ii) Cu(I) and cystine are generated in the reaction; and (iii) there is a positive correlation between the albumin copper content and the reaction rate of oxidation. The role of this redox-cycling mechanism seems to be less relevant in the case of Hcys, since oxidation of this aminothiol is scantly sensitive to copper catalysis. Thus albumin has a much stronger prooxidant effect upon Cys than on Hcys. This assumption is in concordance with the results of Sengupta et al. [5], who reported that albumin mediates the oxidation of Hcys to its disulfide form mainly by thiol disulfide exchange, and to a lesser extent by oxidative processes catalyzed by albumin-bound copper. The in vivo source of the oxidative equivalents driving Cys oxidation has not been identified. Sengupta et al. in in vitro studies [5] showed copper bound to ceruloplasmin to be a highly efficient catalyst for Cys oxidation, and proposed that cystine is formed in circulation primarily by the oxidative catalytic activity of ceruloplasmin. We have found in this study that copper bound to albumin is also an effective catalyst. It must be considered that the concentration of catalytically active Cu(II) afforded by albumin and ceruloplasmin is similar. Assuming albumin and ceruloplasmin plasmatic concentrations of 750 and 2.3 lM, respectively, and an occupancy of 0.3% of the copper binding site in albumin [3], the plasmatic concentration of the copper carried by albumin would be 2.2 lM, i.e., of the same order of that carried by ceruloplasmin, since while the latter binds seven copper atoms, only one is catalytically active in oxidation reactions [26]. This distribution of active copper in equal proportions among ceruloplasmin and albumin only occurs in the postabsorptive period. In the postprandial period albumin is highly enriched in copper, as it functions as a short-lived metabolic compartment for adsorbed copper in its transport from the gastrointestinal tract to the liver [27,28]. In humans, a significant increase in plasma cystine without a concomitant increase in Cys concentration has been reported in the postprandial state [8]. In the postabsorptive state, exportation of amino acids from peripheral tissues is associated with the appearance of cystine but not of Cys [8]. These two observations indicate that Cys from dietary proteins or endogenous pro-
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tein catabolism rapidly oxidizes in plasma, in fact, Cys presents an exceptionally low plasma concentration in comparison with other protein-forming amino acids [29]. These physiological data are consistent with our results, and support the hypothesis that albumin plays a critical role in Cys oxidation through the redox cycling activity of its bound copper atom, mainly in the postprandial period when portal blood is enriched in Cys generated from protein digestion and in copper bound to albumin. The mechanism of Cys oxidation proposed in this work would help explain two aspects of albumin metabolism, which to date have been poorly defined. One aspect would be the formation of nonmercaptalbumin in the circulation, and the other the signaling pathway for albumin degradation. Although albumin is secreted by the liver with its thiol group fully reduced, circulating albumin has 60% of the thiol groups forming a disulfide bond with Cys (nonmercaptalbumin). According to our scheme, cystine generated during Cys oxidation could supply the oxidative equivalents driving oxidation of the albumin thiol group. In fact, this reaction based on thiol disulfide exchange has been used for years for preparing nonmercaptalbumin [30]. Copper catalyzed oxidation of Cys leads to the generation of several reactive oxygen species in a complex process with two-phase kinetics [22]. Modification of secondary and tertiary structures and increased proteolytic susceptibility have been described in albumin exposed to oxygen radicals [31,32]. These radicals could damage albumin in the surroundings of the copper-binding site due to their high reactivity, and the resulting modifications could constitute the signal for selecting an albumin molecule for degradation via binding to scavenger receptors and posterior uptake into endocytic vesicles [3]. Acknowledgment The author thanks Nuria Martin for technical assistance. References [1] J. Masuoka, J. Hegenauer, B.R. Van Dyke, P. Saltman, J. Biol. Chem. 268 (1993) 21533–21537. [2] A. Zgirski, E. Frieden, J. Inorg. Biochem. 39 (1990) 137–148. [3] T. Peters Jr., All about Albumin. Biochemistry, Genetics and Medical Applications, Academic Press, San Diego, 1996. [4] G.J. Quinlan, C. Coudray, A. Hubbard, J.M.C. Gutteridge, J. Pharm. Sci. 81 (1992) 611–614. [5] S. Sengupta, C. Wehbe, A.K. Majors, M.E. Ketterer, P.M. DiBello, D.W. Jacobsen, J. Biol. Chem 276 (2001) 46896–46904. [6] M.A. Mansoor, C. Bergmark, A.M. Svardal, P.E. Lønning, P.M. Ueland, Arterioscler. Thromb. Vasc. Biol. 15 (1995) 232–240. [7] P.M. Ueland, M.A. Mansoor, A.B. Guttormsen, F. Mu¨ller, P. Aukrust, H. Refsum, A.M. Svardal, J. Nutr. 126 (Suppl. 4) (1996) 1281S–1284S.
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[8] W. Dro¨ge, Exp. Gerontol. 37 (2002) 1333–1345. [9] S. Sengupta, H. Chen, T. Togawa, P.M. DiBello, A.K. Majors, B. Bu¨dy, M.E. Ketterer, D.W. Jacobsen, J. Biol. Chem. 276 (2001) 30111–30117. [10] M. Gabaldon, Int. J. Biol. Macromol. 30 (2002) 259–267. [11] C.K. Riener, G. Kada, H.J. Gruber, Anal. Bioanal. Chem. 373 (2002) 266–276. [12] A.R. White, X. Huang, M.F. Jobling, C.J. Barrow, K. Beyreuther, C.L. Masters, A.I. Bush, R. Cappai, J. Neurochem. 76 (2001) 1509–1520. [13] A.J. Brenner, E.D. Harris, Anal. Biochem. 226 (1995) 80–84. [14] A.I. Ivanov, J.A. Parkinson, E. Cossins, J. Woodrow, P.J. Sadler, J. Biol. Inorg. Chem. 5 (2000) 102–109. [15] M. Le, G.E. Means, Anal. Biochem. 229 (1995) 264–271. [16] J.S. Stamler, A. Slivka, Nutr. Rev. 54 (1996) 1–30. [17] C. Perugini, M. Seccia, E. Albano, G. Bellomo, Biochim. Biophys. Acta 1347 (1997) 191–198. [18] L.M. Sayre, Science 274 (1996) 1933–1934. [19] M.D. Apostolova, P.R. Bontchev, B.B. Ivanova, W.R. Rusell, D.R. Mehandjiev, J.H. Beattie, C.K. Nachev, J. Inorg. Biochem 95 (2003) 321–333.
[20] J. Masuoka, P. Saltman, J. Biol. Chem. 269 (1994) 25557–25561. [21] R.F. Chen, J. Biol. Chem. 242 (1967) 173–181. [22] A.V. Kachur, C.J. Koch, J.E. Biaglow, Free Radic. Res. 31 (1999) 23–34. [23] B. Pfanzagl, F. Tribl, E. Koller, T. Mo¨slinger, Atherosclerosis 168 (2003) 39–48. [24] M. Exner, M. Hermann, R. Hofbauer, B. Hartmann, S. Kapiotis, B. Gmeiner, FEBS Lett. 531 (2002) 402–406. [25] E. Nakano, M.P. Williamson, N.H. Williams, H.J. Powers, Biochim. Biophys. Acta 1688 (2004) 33–42. [26] C.K. Mukhopadhyay, B. Mazumder, P.F. Lindley, P.L. Fox, Proc. Natl. Acad. Sci. USA 94 (1997) 11546–11551. [27] D.T. Gordon, A.S. Leinart, R.J. Cousins, Am. J. Physiol. 252 (1987) E327–E333. [28] M.C. Linder, L. Wooten, P. Cerveza, S. Cotton, R. Shulze, N. Lomeli, Am. J. Clin. Nutr. 67 (Suppl) (1998) 965S–971S. [29] A.J. Cooper, Annu. Rev. Biochem. 52 (1983) 187–222. [30] T.E. Isles, P.C. Jocelyn, Biochem. J. 88 (1963) 84–88. [31] K.J.A. Davies, M.E. Delsignore, J. Biol. Chem. 262 (1987) 9908– 9913. [32] G. Marx, M. Chevion, Biochem. J. 236 (1985) 397–400.
Oxidation of cysteine and homocysteine by bovine albumin Marı´a Gabaldon* Unidad de Histoquimia, Centro de Investigacio´n, Hospital La Fe, Avenida Campanar, 21, 46009 Valencia, Spain Received 9 June 2004, and in revised form 27 August 2004
Abstract The autooxidation of cysteine and homocysteine to their disulfide forms was determined by measuring the time course of thiol groups disappearance. We found the oxidative chemistry of cysteine and homocysteine to be quite different. In the absence of added Cu(II), cysteine autooxidized at a slower rate than homocysteine, though in its presence cysteine oxidation was much faster, homocysteine being found to be a poor responder to copper catalysis. Albumin speeded up the spontaneous oxidation of both aminothiols, the reaction being faster with cysteine than with homocysteine. The copper content of different albumins was found to be highly variable, ranging from 12.75 to 0.64 lg Cu(II)/g albumin. We propose that copper bound to albumin possesses redox cycling activity to perform cysteine oxidation since: (i) copper elimination by copper chelators markedly reduces oxidation; and (ii) a positive correlation exists between the albumin copper content and the oxidation reaction rate. 2004 Elsevier Inc. All rights reserved. Keywords: Albumin; Mercaptalbumin; Cysteine oxidation; Homocysteine oxidation; Copper chelators
Albumin is a single chain multidomain protein with 17 disulfide bonds and one free thiol at Cys34, which partially forms mixed disulfides with cysteine (Cys)1 and glutathione. Albumin has a strong binding site for Cu(II) (Ka approaching 1012 M1) [1,2] at the N-terminal site involving the amino terminal nitrogen, the next two peptide nitrogens and the histidyl imidazole nitrogen forming a square planar configuration [3]. The content of Cu(II) in commercial human albumin is highly variable, which suggests that problems of metal contamination or losses arise during manufacturing processes [4]. The copper content described in a recent work was 3.87 lmol Cu(II)/mmol albumin, which indicates *
Fax: +34 6 1973018. E-mail address: [email protected]. 1 Abbreviations used: Cys, cysteine; Hcys, homocysteine; DPDS, 4,4 0 -dipyridyl disulfide; DEPC, diethyl pyrocarbonate; DTT, DL dithiothreitol; EDTA, ethylenediaminetetraacetic acid, disodium salt; DTPA, diethylenetriaminepentaacetic acid; TCA, trichloroacetic acid; Tris, Tris-(hydroxymethyl)aminomethane; BDS, bathocuproinedisulfonic acid, disodium salt; BCA, bicinchoninic acid, disodium salt; Hepes, N-(2-hydroxyethyl)piperazine-N 0 -(2-ethanesulfonic acid). 0003-9861/$ - see front matter 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2004.08.028
that only 0.4% of the albumin molecules carry bound copper [5]. The present study was stimulated by the observation that when preparing aged albumin the –SH groups of Cys and homocysteine (Hcys) completely disappeared after 24 h of incubation with albumin at pH 8.9 and low ionic strength. The fact that the complete oxidation of Cys also occurred when increasing the Cys/albumin molar ratio from 5 to 20 pointed to a catalytic phenomenon where a metal with redox cycling activity, most probably copper, catalyzed Cys oxidation. This observation led us to hypothesize that, under physiological conditions, in the relatively oxidative environment of blood, albumin could play a critical role in the oxidation of Cys and Hcys to their disulfide forms through the catalytic action of the minimal amounts of Cu(II) that it binds and transports. In fact, aminothiols circulate mainly as oxidized forms and only 0.5% of total Hcys and 5% of total Cys are found in the reduced form [6]. Aminothiols in human plasma exist in reduced and free and albumin-bound oxidized forms interacting with each other through redox and disulfide exchange
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reactions. The aminothiols together with the free –SH group of albumin comprise a dynamic system referred to as the redox thiol status and represent the balance between prooxidant and antioxidant processes in plasma [7]. This balance may modulate redox-sensitive signaling pathways and suffers a marked prooxidative shift with aging and in cardiovascular and degenerative diseases [7,8]. The regulation of these redox systems in plasma has been little studied. Recently, Sengupta et al. [5,9] proposed that in circulation, albumin mediates the oxidation of Hcys by thiol disulfide exchange, whereas ceruloplasmin drives Cys oxidation by copper-dependent oxidative mechanisms. The basic question posed in this study is whether the copper bound to albumin has redox-cycling activity to perform the oxidation of aminothiols. We first studied the autooxidation of Cys and Hcys in the absence of albumin, and the response of this reaction to copper catalysis. We found that despite the structural similarity, the chemical behavior was quite different, the oxidation of Hcys being much faster and much less sensitive to copper catalysis than that of Cys. To know whether the effect we first observed was a general phenomenon we studied albumins with different compositions and purities to exclude the possible effect of copper-rich contaminant globulins. All studied albumins showed a prooxidant effect on Cys and this effect was related to the copper content of the albumin. Finally, in order to demonstrate that copper bound to albumin was involved in Cys oxidation, experiments were performed after removing copper with different types of copper chelators. Results indicate that albumin may play a crucial role in Cys oxidation by a copper-catalyzed oxidative process and that copper-dependent Hcys oxidation by albumin is a minor pathway in the overall conversion to disulfide forms. As proposed by Sengupta et al. [5], a thiol disulfide exchange with the disulfide bond at Cys34 of albumin seems to be a more plausible mechanism for Hcys oxidation.
Materials and methods Chemicals The following bovine serum albumins were supplied by Sigma Chemical:A-4503 lot 050K0891 remaining most globulins; A-0281 essentially fatty acid and globulin free; A-7638 essentially globulin free; A-7511 essentially fatty acid free; and A-3912 remaining most globulins and obtained by heat shock. The following bovine serum albumins were supplied by ICN Biomedicals: 194774 essentially globulin free and 810012 crystalline. Bicinchoninic acid disodium salt, bathocuproinedisulfonic acid disodium salt, L -cysteine free base (C-7352), DL -homocysteine, L -histidine (H-8000), diethylenetri-
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aminepentaacetic acid (D-6518), DL -dithiothreitol (D0632), iodoacetamide (I-6125), L -ascorbic acid (A-5960), diethyl pyrocarbonate, and trichloroacetic acid (T9159) were supplied by Sigma Chemical Co. Copper atomic spectroscopy standard solution and 4, 4 0 -dipyridyl disulfide (DPDS; Aldrithiol-4) were supplied by Fluka and Aldrich, respectively. Solutions were prepared in deionized water obtained with an Elix water purification system from Millipore (10–15 MX cm resistivity). Buffered albumin solutions were prepared as previously described [10]. Determination of thiol groups Thiol groups were determined with DPDS as described by Riener et al. [11] with some modifications. The DPDS reagent is a 4 mM DPDS solution in 12 mM HCl and was prepared by dissolving 45 mg of DPDS in 1.5 ml of 0.4 M HCl before completing with H2O to 50 ml. Aliquots were immediately frozen at 20 C and thawed portions were used within 1 h. Reaction was performed by mixing 1.2 ml of the study sample in 0.1 M sodium phosphate buffer pH 6 with 50 ll of the DPDS reagent. Albumin concentration for thiol group determination must be between 0.5 and 5 mg/ml and aminothiol concentration between 4 and 40 lM. A reagent blank with buffer instead of study sample and a sample blank with buffer instead of DPDS were processed in the same way. Absorbances at 324 nm were measured against H2O after 10 min. The thiol group reacts with DPDS to produce an equivalent of 4-thiopyridone, which has a molar absorptivity of 21,400 M1 cm1 at 324 nm. The concentration of 4-thiopyridone was calculated from the absorbance at 324 nm after subtracting the absorbances of the two blanks. Determinations were performed in duplicate. Size exclusion chromatography In some experiments albumin was incubated with low molecular weight reagents, which after reaction must be completely removed by gel filtration. The separation of albumin from low molecular weight products was performed by passing solutions through a Hi Prep 26/10 desalting column (Amersham Biosciences) equilibrated with H2O. A 10-ml sample of A-4503 albumin filtered through 0.45 lm was applied at 5 ml/min followed by 1 ml washing with H2O; elution was performed with H2O at 2.5 ml/min and the albumin fraction was collected between 2 and 8 min after the start of elution. Given that the molar concentration of the reagents considerably exceeded that of albumin, controls were performed to make sure that the eluate fraction with albumin was free from low molecular weight products. The pattern of histidine elution was obtained from a 45-mM solution by monitoring the conductivity in
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eluates or by histidine determination with diethyl pyrocarbonate (DEPC). In this latter case 600 ll of eluate were mixed with 580 ll of 0.1 M sodium phosphate buffer pH 7.5 and with 20 ll of 91 mM DEPC in acetonitrile prepared as indicated in [10]. Readings were taken 15 min later at 240 nm. The pattern of DL -dithiothreitol (DTT) elution was obtained by determination of the thiol groups in eluates after applying a 1.8 mM DTT solution. Iodoacetamide, ethylenediaminetetraacetic acid (EDTA), and diethylenetriaminepentaacetic acid (DTPA) could be determined by monitoring the conductivity of the effluents. All the determinations showed the fraction collected between 2 and 8 min under the described conditions to contain albumin without the presence of low molecular weight substances.
thione. A 72 mg/ml A-4503 albumin solution in 0.3 M NaCl–0.1 M sodium phosphate buffer (final pH 6.85) (10 ml) was mixed with a 10.8 mM DTT solution in 0.3 M NaCl–0.1 M sodium phosphate buffer pH 6.75 (2 ml), keeping the final pH between 6.76 and 6.84. The reaction mixture was filtered through a 0.45-lm filter, and after 25 min of incubation a 10 ml sample was applied to a Hi Prep 26/10 desalting column under the conditions previously described. The collected albumin fraction was analyzed for thiol groups after diluting 1/ 20 with 0.1 M sodium phosphate buffer pH 6. The – SH/albumin molar ratio was found to be 1.00 ± 0.05 (mean ± SD, n = 4). The composition of mercaptalbumin determined by gel filtration chromatography was the same as the starting albumin (86.1% monomer, 11.8% dimer, and 2.1% polymers).
Autooxidation of aminothiols Blocking of albumin –SH groups The reaction mixture at pH 7.4 contained: aminothiol, 1.5 mM; Tris–HCl buffer pH 7.5, 10 mM; and NaCl, 150 mM. The reaction mixture at pH 8.9 was: aminothiol, 1.5 mM; and Tris–HCl buffer pH 9, 50 mM. The 25 mM starting aminothiol solutions were freshly prepared. Incubations were performed for 24 h at room temperature in the dark. Samples were taken at 2, 4, 8, and 24 h, diluted (1/40 to 1/10) with 0.1 M sodium phosphate buffer pH 6, and thiol groups were determined by DPDS analysis. Values at t = 0 were obtained by diluting the starting aminothiol solution with H2O to 1.5 mM. After 1/40 dilution with 0.1 M sodium phosphate buffer pH 6 and 4 min of incubation, samples were taken for thiol determination. Oxidation of aminothiols by albumin The reaction mixture contained: albumin, 300 lM; NaCl, 150 mM; and aminothiol, 1.5 mM; pH adjusted to 7.4 with 0.2 M Tris. Aminothiol solutions were freshly prepared. Albumin solution was filtered through a 0.45-lm filter. Incubations, values at t = 0, and DPDS analysis were performed as indicated in the previous section. Total thiol groups in the reaction mixture were the sum of the added thiol groups (1500 lM) and the thiol groups carried by the albumin. The –SH/albumin molar ratio of albumins ranged from 0.40 to 0.55, and was calculated by determination of the thiol groups in a 60 lM albumin solution in 0.1 M sodium phosphate buffer pH 6. The concentration (lM) of the albumin thiol groups was obtained by multiplying 300 by the –SH/albumin molar ratio. Preparation of mercaptalbumin Mercaptalbumin has the thiol group at Cys34 fully reduced whereas normal albumin has close to 50% of this thiol group forming disulfide bonds with Cys and gluta-
The free –SH group of albumin was blocked with iodoacetamide. The final concentrations in the reaction mixture were: albumin, 900 lM; iodoacetamide, 4.5 mM; NaCl, 50 mM; final pH adjusted with 0.2 N NaOH, 7.5. Incubation was carried out for 1 h in the dark, and the solution was passed through a Hi Prep 26/10 desalting column under the conditions previously described. The collected albumin fraction was negative for thiol groups. Copper calibration curve A 1 mM Cu(NO3)2–6 mM glycine stock solution was prepared from a copper atomic spectroscopy standard solution. Copper–glycine stock solutions were used to prevent the formation of metal-hydroxy and metal-oxy polymers in neutral media [12]. Copper(II) was determined after reduction by ascorbic acid by spectrophotometric analysis at 358 nm of the Cu(I) complex formed with bicinchoninic acid (BCA). As BCA from different suppliers showed important differences in capacity to complex Cu(I), before starting determinations BCA must be assayed by adding increasing amounts to a fixed copper concentration covering a BCA/Cu(I) molar ratio range from 2 to 3 at 0.2 intervals, until absorbances are near constant. As the molar absorptivity of the BCA– Cu(I) complex is approximately 23 times greater than that of BCA, absorbance increases notably when increasing BCA concentration if this is defective, and very discretely when in excess. In our case the BCA/ Cu(I) molar ratio for complete copper complexation was 2.3. Calibration curves were prepared under the conditions described for the analysis of copper in albumin (section below), except that BCA solution in 0.6 M sodium Hepes–0.15 M aOH was used instead of 0.6 M Hepes buffer pH 8. In all cases the final pH was 6.8–7.2, and the BCA/Cu(I) molar ratio was 2.3. Extinc-
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tion coefficients were calculated from calibration curves after subtracting absorbances of blanks without copper and BCA from those of the study samples. Under these conditions, the extinction coefficient of the BCA–Cu(I) complex at 358 nm was 45.37 ± 1.62 mM1 cm1 (mean ± SD, n = 23), and that of BCA was found to be 1.94 ± 0.28 mM1 cm1 (mean ± SD, n = 23). Copper determination in albumin The copper bound to albumin was determined by a modification of the technique described by Brenner and Harris [13], based on the separation of copper(II) in strong acid media, reduction to Cu(I) with ascorbic acid, and formation of a complex with BCA, which is then spectrophotometrically assayed. An albumin solution was prepared by mixing 1.2 g of albumin and 3.6 ml of H2O in a 10-ml vial with magnetic stirring. After 1/160 dilution, readings at 280 nm were taken and albumin concentration was adjusted to 3.5 mM (extinction coefficient of albumin at 280 nm, 65,500 M1 cm1). Albumin solution was mixed with 30% trichloroacetic acid (TCA) in a 3/1 ratio v/v. The suspension was stirred in a 10-ml vial for 2 h and centrifuged two times at 5000 rpm for 20 min. The supernatant obtained under these conditions represented 60% of the initial suspension volume. Copper determination was performed by mixing in the order stated: 0.60 ml of supernatant, 0.12 ml of freshly prepared 2 mM ascorbic acid, and 0.48 ml of 60 or 100 lM BCA in 0.6 M Hepes buffer pH 8. A blank without BCA was processed in the same way. Readings were taken at 358 nm against H2O, and the absorbance of the blank was subtracted from that of the study sample. As BCA forms a 2:1 complex with Cu(I), BCA concentration must be greater than double that of Cu(I) but not too high to minimize interferences produced by excess BCA, which also absorbs, though discretely, at 358 nm. To conciliate these two requirements, BCA concentration has been adapted to that of Cu(I) in the following way: beginning with a 60 lM BCA concentration, absorbance of the study sample minus the blank must be <0.5; if absorbance >0.5, then a 100 lM BCA concentration must be used. When absorbance is >0.88 with this last concentration, the supernatant must be diluted 1/2 with 4.5% TCA. This scheme covers a range of copper concentrations in the supernatant from 6 to 80 lM. The absorbance of the reaction mixture is the sum of the absorbances of the BCA–Cu(I) complex and excess BCA. Copper concentration in the reaction mixture (lM) was calculated as =(a b * c)/(d 2b) where a = absorbance of the study sample minus that of the blank; b = lM absorptivity of BCA (1.94 · 103); c = initial BCA concentration in the reaction mixture (lM); and d = lM absorptivity of the BCA–Cu(I) complex (45.37 · 103). Copper concentration in the starting 3.5 mM albumin
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solution was calculated by multiplying copper concentration in the reaction mixture by 2 and dividing by 0.75/0.6. Determinations were performed in duplicate. Copper reduction assay in the absence of albumin Aminothiols in the presence of added Cu(II) oxidize and produce Cu(I). Hcys oxidation by copper was studied at different molar ratios by monitoring the formation of Cu(I) with bathocuproinedisulfonic acid (BDS) or BCA. The reaction mixture contained: Cu(II) (40 lM Cu(NO3)2–240 lM glycine), 40 lM; sodium phosphate buffer pH 7.4, 20 mM; NaCl, 150 mM; BDS, 120 lM; and freshly prepared Hcys, 400 or 40 lM for a Hcys/ Cu(II) molar ratio of 10 or 1, respectively. When using BCA, final concentrations of BCA and Hepes buffer pH 7.4 were 92 lM and 20 mM, respectively. Reaction was started by addition of Hcys. Copper reduction was monitored by registering the absorbance of the complex BDS–Cu(I) at 480 nm, considering an absorptivity of 12,540 M1 cm1 [14]. When using BCA, Cu(I) in the reaction mixture was determined by absorbance at 358 nm after applying the formula described in the previous section. To correlate formation of Cu(I) and disappearance of thiol groups at different aminothiol/Cu(II) molar ratios, incubation mixtures were prepared containing: Cu(II) (40 lM Cu (NO3)2–240 lM glycine), 40 lM; sodium phosphate buffer pH 7.4, 20 mM; NaCl, 150 mM; and freshly prepared aminothiol, 400 or 40 lM for a aminothiol/Cu(II) molar ratio of 10 or 1, respectively. Reaction was started by addition of the aminothiol, and after 5 min samples were taken for DPDS analysis. When using an aminothiol/Cu(II) molar ratio of 10, samples were diluted 1/10 with 0.1 M sodium phosphate buffer pH 7.4, while the samples were used undiluted when the molar ratio was 1. Values at t = 0 were obtained by diluting the starting aminothiol solution with 20 mM sodium phosphate buffer pH 7.4 to 40 lM. Copper reduction assay in the presence of albumin Since techniques for Cu(I) determination are not sufficiently sensitive to detect the low amount of copper bound to albumin, a copper enriched albumin with 50% occupation of its Cu(II) binding site was prepared by the addition of exogenous Cu(II). Binding of Cu(II) to albumin was performed by incubating for 50 min equal volumes of 150 lM Cu(II) (150 lM Cu(NO3)2 and 900 lM glycine) and 300 lM of A-4503 albumin in 0.1 M sodium phosphate buffer, final pH 7.4. For assessing complete copper binding to albumin, an aliquot was filtered through a 30 K Omega-type Filtron membrane in an Amicon 50 ml ultrafiltration cell, and the Cu(II) content was analyzed in the filtrate. Similar incubation with buffer instead of albumin was carried
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out. The reaction mixture contained the following: NaCl, 150 mM; albumin, 100 lM; Cu(II) (50 lM Cu(NO3)2–300 lM glycine), 50 lM; BDS, 150 lM; and freshly prepared aminothiol, 500 lM. Reaction was started by addition of aminothiol. Copper reduction was monitored by registering the absorbance of the complex BDS–Cu(I) at 480 nm, considering an absorptivity of 12,540 M1 cm1. Similar reaction mixtures were prepared with the filtrate obtained by passing the Cu(II) enriched albumin through a 30 K membrane. The net absorbances were calculated by deducting the absorbances of the blanks prepared without thiol from the absorbances of the study samples. Disulfide reduction assay To demonstrate that aminothiols in the presence of albumin were oxidized to their disulfide forms, disulfide bonds were reduced by DTT, and the formed thiol groups were determined after complexation of excess DTT by arsenite [15]. Albumin (300 lM) was incubated with Cys or Hcys (1500 lM) in 150 mM NaCl, pH 7.4 for 24 h and was then filtered through a Filtron membrane 30 K Omega-type in an Amicon 50 ml ultrafiltration cell. Filtrate was diluted 1/2, adjusting the final pH to 8 with Tris–HCl buffer pH 8. This solution was mixed with an equal volume of 3.8 mM DTT–1 mM EDTA in 50 mM Tris–HCl buffer pH 8, and incubated for 70 min under nitrogen. Reaction mixture was diluted 1/10 with 3.8 mM sodium arsenite in 0.1 M sodium phosphate buffer pH 6 and incubated for 8 min. Thiol groups were immediately determined by DPDS reaction. A blank with 20 mM Tris–HCl buffer pH 8 instead of filtrate was processed in the same way, and its absorbance subtracted from that of the study samples. Analytical procedures Fractionation of albumin in monomeric and polymeric forms was performed by gel filtration using Superdex 200 prep grade on a HiLoad 16/60 column (Amersham Biosciences). The column was loaded with 1 ml 20 mg/ml albumin solution and eluted with 0.15 M NaCl in 10 mM sodium phosphate buffer pH 7.4 at a pumped flow rate of 0.8 ml/min.
Results Autooxidation of aminothiols Fig. 1A shows the effect of pH on the time course of aminothiol oxidation measured by the loss of –SH groups in mixtures without added copper. Under physiological conditions (pH 7.4 and 150 mM NaCl) the rate of autooxidation is faster for Hcys than for Cys. This
Fig. 1. Autooxidation of aminothiols. Time course of –SH group disappearance. (A) Effect of pH. The reaction mixture at pH 7.4 contained: aminothiol, 1.5 mM; Tris–HCl buffer pH 7.5, 10 mM; and NaCl, 150 mM. The reaction mixture at pH 8.9 was: aminothiol, 1.5 mM; and Tris–HCl buffer pH 9, 50 mM; Cys pH 7.4 (——); Cys pH 8.9 (—j—); Hcys pH 7.4 (—m—); Hcys pH 8.9 (—d—). (B) Effect of Cu(II). The reaction mixture contained: aminothiol, 1.5 mM; Tris–HCl buffer pH 7.5, 10 mM; NaCl, 150 mM; and Cu(II) (0.61 lM Cu(NO3)2–3.67 lM glycine), 0.61 lM. Final pH, 7.4; Cys (——); Cys + Cu (II) (—d—); Hcys (—m—); Hcys + Cu(II) (—j—). The amount of remaining –SH groups was determined by DPDS analysis, as described in Materials and methods. Each point represents the mean from five independent experiments. The coefficient of variation did not exceed 20%.
may be explained taking into account that Hcys is a stronger nucleophile than Cys, and is thus more reactive [16]. The rise in pH to 8.9 practically does not modify the rate of Cys oxidation but drastically increases that of Hcys due to the lower pKa of the thiol group of the former [9]. Fig. 1B shows the time course of the oxidation of aminothiols in the presence of added Cu(II) measured by the loss of –SH groups. A low copper concentration (0.61 lM) has been used in order to compare the effect of free Cu(II) added to the reaction mixture with the effect of Cu(II) bound to commercial albumin described in the following section in which the same final copper concentration was obtained from a 300 lM A-4503 albumin solution (Table 1). Copper(II) was much more efficient in catalyzing Cys oxidation than Hcys oxidation. At 2 h, Cys lost 1% of its –SH
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Table 1 Cysteine oxidation and albumin copper content Albumin
Loss of –SH groups (%)a
Copper contentb lgCu(II)/g albumin
2h
4h
8h
24 h
A-4503 810012 A-3912
46.6 94.9 83.3
74.8 97.9 95.9
93.9 99.2 98.6
98.4 99.7 99.5
1.95 12.75 2.33
Globulin free A-7638 194774
40.1 89.4
64.1 96.4
88.6 98.5
98.0 99.6
1.44 4.42
Fatty acid free A-7511 A-0281
38.9 15.6
62.3 31.6
86.8 52.4
96.7 90.8
0.73 0.64
a The reaction mixture contained: albumin, 300 lM; NaCl, 150 mM; and Cys, 1.5 mM; pH adjusted to 7.4 with 0.2 M Tris. The amount of the remaining –SH groups was determined by DPDS analysis, as described in the ‘‘Materials and methods’’ section. Each value represents the mean from three independent experiments. The coefficient of variation did not exceed 15%. b Each value represents the mean from three independent determinations. The coefficient of variation did not exceed 10%.
groups in the absence and 94% in the presence of Cu(II), whereas the losses in –SH groups for Hcys at the same time were 7 and 17%, respectively. Oxidation of aminothiols in the presence of Cu(II) can be determined by measuring the disappearance of thiol groups or the formation of Cu(I) with chelating reagents such as BCA or BDS. The time course of Hcys oxidation was different depending on the reagent used for determination of the formed Cu(I) (Fig. 2). At a Hcys/Cu(II) molar ratio of 10, the kinetics with BCA was characterized by a lag-phase of 15 min followed by a period where Cu(I) was formed at a slow rate. The reaction was dependent on the buffer used, being
Fig. 2. Autooxidation of Hcys by Cu(II). Time course of Cu(I) formation. The reaction mixture contained: Cu(II) (40 lM Cu (NO3)2– 240 lM glycine), 40 lM; NaCl, 150 mM; BCA or BDS, 92 lM or 120 lM, respectively; Hepes buffer or sodium phosphate buffer pH 7.4, 20 mM; and Hcys, 40 or 400 lM. Reaction was started by addition of Hcys. Numbers in parentheses indicate the Hcys/Cu(II) molar ratio. BCA (1) (—m—); BCA (10) (- -m- -); BCA (10) with phosphate buffer (—d—); BDS (1) (——); BDS (10) (—j—). The amount of formed Cu(I) was determined by absorbance at 480 and 358 nm when using BDS and BCA, respectively, as described in Materials and methods. Each point represents the mean from three independent experiments.
faster with phosphate than with Hepes buffer at the same pH of 7.4. When using BDS, the kinetics was completely different, the reduction of Cu(II) being almost instantaneous. Other studies described a marked stimulation of copper-induced oxidation of LDL by BDS [17]. A possible explanation for these observations is that BDS also binds Cu(II), and this complex is a stronger oxidant than Cu(II) [14,18]. Thus, BDS must be used with caution to monitor Cu(II) reduction, because thiol oxidation measured in its presence could generate methodological artifacts. When Cys was incubated with Cu(II) at molar ratios of 1 and 10, the reduction of Cu(II) was almost instantaneous regardless of whether BDS or BCA was used for determination of the formed Cu(I) (data not shown). An unexpected finding of this study was the influence of the Hcys/Cu (II) molar ratio on the time course of the reaction when using BCA. At a Hcys/Cu(II) molar ratio of 1, copper reduction was almost instantaneous, whereas the reaction rate was much slower at a Hcys/ Cu (II) of 10 (Fig. 2). The same results were obtained when Hcys oxidation was measured by disappearance of thiol groups. Fig. 3 shows the oxidation of aminothiols after 5 min of incubation with Cu(II), expressed as the disappearance of thiol groups or the appearance of Cu (I). Results indicate that when using BCA for determination of Cu(I), both criteria were concordant except for a minor imbalance found at Hcys/Cu(II) molar ratio of 10 that we are unable to account for. It has recently been reported [19] that the interaction between Cu(II) and Hcys at molar ratio 1:1 leads to the formation of a blue complex with Cu (II), and that in the presence of excess ligand the complex is yellow with copper in the reduced state. The differences we have observed at Hcys/Cu(II) molar ratios of 1 and 10 may be related to the oxidative capacity of the different Hcys/copper complexes formed.
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Fig. 3. Autooxidation of aminothiols by Cu(II). –SH group disappearance and Cu(I) formation at 5 min For determination of the remaining –SH groups, the reaction mixture contained: Cu(II) (40 lM Cu(NO3)2–240 lM glycine), 40 lM; NaCl, 150 mM; sodium phosphate buffer pH 7.4, 20 mM; and aminothiol, 40 or 400 lM. Reaction was started by the addition of aminothiol. For determination of the Cu(I) formed, the reaction mixture contained: Cu(II) (40 lM Cu(NO3)2– 240 lM glycine), 40 lM; NaCl, 150 mM; BCA, 92 lM; Hepes buffer, pH 7.4, 20 mM; and aminothiol, 40 or 400 lM. Reaction was started by the addition of aminothiol. Numbers indicate the aminothiol/Cu(II) molar ratio. Dotted bars, disappearance of –SH groups. Dashed bars, formation of Cu (I). The amount of formed Cu(I) was determined with BCA by measuring the absorbance at 358 nm, and the amount of remaining –SH groups was determined by DPDS analysis, as described in Materials and methods. Values plotted are the mean from three independent experiments.
Oxidation of aminothiols by albumin Fig. 4 shows the time course of the oxidation of aminothiols by albumin at pH 7.4, measured by the loss of thiol groups. Albumin induces a loss of –SH groups in both aminothiols, the reaction rate being faster for
Fig. 4. Oxidation of aminothiols by albumin. Time course of –SH group disappearance. The reaction mixture contained: albumin (A-4503), 300 lM; NaCl, 150 mM; and aminothiol, 1.5 mM; pH adjusted to 7.4 with 0.2 M Tris; Cys (——); Cys + albumin (—d—); Hcys (—m—); Hcys + albumin (—j—). The amount of the remaining –SH groups was determined by DPDS analysis, as described in Materials and methods. Each point represents the mean from five independent experiments. The coefficient of variation did not exceed 20%.
Fig. 5. Aminothiol oxidation by albumin. Time course of Cu(I) formation. The reaction mixture contained: albumin (A-4503), 100 lM; Cu(II) (50 lM Cu (NO3)2–300 lM glycine), 50 lM; NaCl, 150 mM; aminothiol, 500 lM; BDS, 150 lM; sodium phosphate buffer, pH 7.4, 20 mM; albumin and Cu(II) were incubated for 50 min before addition of the remaining reagents. Reaction was started by the addition of aminothiol; Cys (—j—); Cys + albumin (——); Hcys (—m—); Hcys + albumin (—d—). The amount of Cu(I) formed was determined by absorbance at 480 nm, as described in Materials and methods. Each point represents the mean from three independent experiments.
Cys than for Hcys. Thiol groups disappearance was due to the formation of disulfide bonds since 95–100% of the initial –SH groups were recovered from the filtrate through a 30 K membrane after DTT reduction and complexing excess DTT by arsenite. To examine whether the copper bound to albumin had redox cycling activity, aminothiols were incubated with a Cu(II) enriched albumin (albumin/Cu(II) molar ratio of 2), and the formed Cu(I) was assayed with BDS. Under the conditions used complete copper binding to albumin was produced, since the filtrate from a 30 K membrane was negative for copper; results thus refer to the effect of copper bound to albumin and not to any residual free copper. Fig. 5 shows the time course of the formation of Cu(I). In the presence of free Cu(II) and in the absence of albumin, the reduction of Cu(II) by aminothiols was nearly instantaneous. In the presence of albumin, the reduction of Cu(II) was delayed with both aminothiols, but was much more faster with Cys than with Hcys. Results indicate that copper bound to albumin has redox cycling activity, and that aminothiols reduce free Cu(II) faster than Cu(II) bound to albumin. To study the role of the albumin thiol group at Cys34 in mediating the oxidation of Cys, we performed incubations with mercaptalbumin and with albumin with the free –SH group blocked with iodoacetamide. As shown in Fig. 6, the time course of the reaction was similar in the presence and in the absence of free thiol groups, which indicates that the thiol group of albumin is not involved in the oxidation of Cys. As albumin A-4503 is a fraction V remaining most globulins there was the possibility that contaminant pro-
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EDTA. When the EDTA–Cu(II) complex was not removed from the incubation mixture, inhibition was not produced, and Cys oxidation at 2 h was 47% greater than that seen with albumin, possibly due to the greater accessibility of Cys to the copper bound to EDTA than to the copper bound to albumin (Fig. 7A). We tried to remove copper from albumin by incubating the latter with a large excess of histidine, since histidine forms chelates with Cu(II). When before beginning reaction albumin was incubated with histidine in a 1/50
Fig. 6. Oxidation of Cys by albumin. Effect of the albumin thiol group. Time course of –SH group disappearance. The reaction mixture contained: albumin A-4503, mercaptalbumin or –SH blocked albumin, 300 lM; NaCl, 150 mM; and Cys, 1.5 mM; pH adjusted to 7.4 with 0.2 M Tris; Cys (——); Cys + albumin (—j—); Cys + mercaptalbumin (—m—); Cys + albumin (–SH blocked) (—d—). The amount of the remaining –SH groups was determined by DPDS analysis, as described in Materials and methods. Each point represents the mean from five independent experiments. The coefficient of variation did not exceed 15%.
teins, mainly ceruloplasmin, played a significant role in the oxidation. Additional studies carried out with globulin-free albumins and with the purest albumins showed that Cys oxidation was produced in the absence of globulins, and that the reaction rate was variable and independent of albumin purity (Table 1). The molecular weight of ceruloplasmin (132 kDa) is similar to that of albumin dimer (133 kDa). Gel filtration completely resolved the monomeric and dimeric forms of albumin, albumin monomer can thus be considered to be free from any contaminant ceruloplasmin. Albumin A-4503 monomer oxidized Cys at a higher rate than unfractionated albumin, the disappearance of thiol groups being 94 and 47% at 2 h for monomer and unfractionated albumin, respectively. This high reactivity indicated that Cys oxidation was due to albumin itself and not to contaminant high molecular weight substances. Oxidation of cysteine by albumin: Effect of chelating agents To determine whether the copper bound to albumin played a role in Cys oxidation, assays were performed in the presence of chelating agents. Incubations in the presence of 1 mM EDTA failed to inhibit Cys oxidation (data not shown). However, when before beginning the reaction albumin was incubated with EDTA in a 1/10 molar ratio at pH 7.4 for 20 h and the EDTA–Cu(II) chelate was removed by gel filtration, the reaction rate was considerably slowed and Cys oxidation was inhibited by 91 and 43% at 2 and 24 h, respectively The same results were obtained when using DTPA instead of
Fig. 7. Oxidation of Cys by albumin. Time course of –SH group disappearance. (A) Effect of chelating agents. The reaction mixture contained: albumin (A-4503), 300 lM; NaCl, 150 mM; and Cys, 1.5 mM; pH adjusted to 7.4 with 0.1 M Tris. Albumin was preincubated with EDTA or DTPA in molar ratio 1/10 for 20 h at pH 7.4 and low molecular weight components removed by gel filtration; Cys (——); Cys + albumin (- -j- -); Cys + albumin incubated with EDTA (—j—); Cys + albumin incubated with EDTA (without desalting) (—d—); Cys + albumin incubated with DTPA (—m—). (B) Other effectors. The reaction mixture contained of albumin (A-4503), 300 lM; NaCl, 150 mM; and Cys, 1.5 mM; pH adjusted to 7.4 with 0.1 M Tris. BDS (100 lM) was added after Cys. Albumin was preincubated with histidine in molar ratio 1/50 at pH 7 for 20 h and low molecular weight components removed by gel filtration. Albumin was preincubated with HNO3 at pH 1.8 for 20 h and passed through a desalting column; Cys (——); Cys + albumin (- -j- -); Cys + albumin incubated with histidine (—m—); –cys + albumin incubated with HNO3 (—j—); Cys + albumin + BDS (—d—). The amount of remaining –SH groups was determined by DPDS analysis as described in Materials and methods. Each point represents the mean from five independent experiments. The coefficient of variation did not exceed 20%.
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molar ratio at pH 7 for 20 h and the histidine–Cu(II) complex was removed by gel filtration, the reaction rate was considerably slower than that of untreated albumin, and Cys oxidation was inhibited by 96 and 75% at 2 and 24 h, respectively (Fig. 7B). An increase in the histidine/ albumin molar ratio to 90 did not modify the results. However, when incubation with histidine was performed at pH 5, the rate of Cys oxidation was increased, reaching intermediate values between the untreated and treated albumin at pH 7 (data not shown). This pH dependency may be explained by the lower copper binding capacity of histidine at pH 5 than at pH 7 [20]. For removing copper from albumin, apart from these metal binding strategies with low molecular weight chelators, we used acid conditions, since binding of copper to albumin increases linearly from pH 4 to 7, and is minimal at lower pH values [20]. When before beginning the reaction albumin was incubated with HNO3 at pH 1.8 for 20 h and passed through a desalting column, the reaction rate was considerably slower than that of untreated albumin and Cys oxidation was inhibited by 91 and 71% at 2 and 24 h, respectively (Fig. 7B). As Cys oxidation was coupled to a continuous copper redox cycling process, we tried to inhibit reaction by avoiding reoxidation of the formed Cu(I) by atmospheric oxygen. BDS was added after Cys to the reaction mixture in order to form a redox inactive complex with Cu(I). Cys oxidation was inhibited by 68 and 44% at 2 and 24 h, respectively. Results indicate that the oxidation of Cys is coupled to the reoxidation of Cu(I) in albumin, as the reaction rate decreases when the availability of Cu(I) for redox cycling is impaired by BDS complexing (Fig. 7B). Albumin copper content As is shown in Table 1, the copper content of different commercial albumins (fraction V, crystalline, fatty acid free, and globulin free) was highly variable, and was not correlated to the presence of globulins. Elimination of globulins was not followed by a decrease in copper content, which indicates that copper is afforded by albumin itself and not by contaminant copper-rich globulins. The purest albumin (810012) had the greatest copper content (12.75 lg Cu(II)/g albumin), representing 1.3% occupancy of the copper-binding site. Contrarily, fatty acid free albumins (A-7511 and A-0281) were the poorest, with 0.73 and 0.64 lg Cu(II)/g albumin, which represents 0.08 and 0.07% of albumin molecules with bound copper, respectively. As indicated by the manufacturer, fatty acid free albumins were obtained by the method of Chen [21], which is based on the removal of fatty acids by adsorption on charcoal in strong acid medium. This condition does not favor copper binding to albumin, and apart from fatty acids the treatment can remove some of the bound copper.
Table 1 shows the time course of thiol group disappearance after incubation of different commercial albumins with Cys. At 2 h there was a positive correlation between albumin copper content and percentage thiol group loss (n = 6, r = 0.86, P < 0.05). Albumin 810012 has been excluded from this calculation, because its reaction rate was much higher than that of the rest of albumins, having lost 80% of the thiol groups at 40 min, and almost 100% at 2 h.
Discussion The oxidation of aminothiols in the presence of copper is a complex phenomenon due to interaction between copper and thiols with the formation of chelates. In the case of Cys it has been suggested that the reaction catalyst is not the free metal but a 1:2 copper–thiol complex, where the close location of the free amino group and thiol group is crucial for thiol oxidation [22]. According to this, formation of the aminothiol–copper complex would be favored in the case of Cys, due to its shorter chain length and this would explain why Hcys despite oxidizing at a greater rate than Cys in the absence of copper, would oxidize at a slower rate in its presence. Our results help explain the apparently contradictory observations of previous reports on the effect of Cu(II) upon aminothiol oxidation. When using a high aminothiol/Cu(II) molar ratio and measuring oxidation by disappearance of the thiol groups, some authors reported Cys oxidation to be much faster than Hcys oxidation, [5,23], whereas the opposite was found by other investigators when aminothiol/Cu(II) was <1 and oxidation was measured by Cu(I) assay [12,24]. In relation to these results, it has been shown that the lag times for oxidation of low density lipoprotein lipids in the presence of Cu(II) vary markedly as a function of Hcys concentration: a Hcys/Cu(II) molar ratio <1 shortens the lag time, while a molar ratio > 1 slows oxidation [25]. All these results indicate that Hcys oxidation in the presence of copper is dependent on their relative concentrations, and this fact must be taken into account when extrapolating data obtained by experimental protocols to the in vivo situation, where the actual molar ratio of the components may be quite different. We found the time course of Cys oxidation to be similar in the presence and in the absence of free –SH groups, which indicates that the albumin thiol group is not involved in Cys oxidation. These results are in discordance with those obtained by Sengupta et al. [5], who reported complete inhibition of the oxidation of both aminothiols when using mercaptalbumin. The procedure followed by these authors to obtain mercaptalbumin (DTT/albumin molar ratio of 5, 45 min incubation, and prolonged elimination of DTT by dialysis) favored
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copper removal from albumin more than our conditions (DTT/albumin molar ratio of 2, 25 min incubation, and rapid DTT elimination by desalting), and, in fact, the mercaptalbumin obtained by these authors had 94% less copper than the starting albumin. The purpose of this study was to demonstrate whether the minimal amount of copper bound to albumin under physiological conditions would be able to catalyze the oxidation of aminothiols by a redox cycling mechanism. Results indicate that copper bound to albumin exhibits a remarkable catalytic redox-cycling activity towards Cys since: (i) elimination of Cu(II) by copper chelators markedly decreases oxidation; (ii) Cu(I) and cystine are generated in the reaction; and (iii) there is a positive correlation between the albumin copper content and the reaction rate of oxidation. The role of this redox-cycling mechanism seems to be less relevant in the case of Hcys, since oxidation of this aminothiol is scantly sensitive to copper catalysis. Thus albumin has a much stronger prooxidant effect upon Cys than on Hcys. This assumption is in concordance with the results of Sengupta et al. [5], who reported that albumin mediates the oxidation of Hcys to its disulfide form mainly by thiol disulfide exchange, and to a lesser extent by oxidative processes catalyzed by albumin-bound copper. The in vivo source of the oxidative equivalents driving Cys oxidation has not been identified. Sengupta et al. in in vitro studies [5] showed copper bound to ceruloplasmin to be a highly efficient catalyst for Cys oxidation, and proposed that cystine is formed in circulation primarily by the oxidative catalytic activity of ceruloplasmin. We have found in this study that copper bound to albumin is also an effective catalyst. It must be considered that the concentration of catalytically active Cu(II) afforded by albumin and ceruloplasmin is similar. Assuming albumin and ceruloplasmin plasmatic concentrations of 750 and 2.3 lM, respectively, and an occupancy of 0.3% of the copper binding site in albumin [3], the plasmatic concentration of the copper carried by albumin would be 2.2 lM, i.e., of the same order of that carried by ceruloplasmin, since while the latter binds seven copper atoms, only one is catalytically active in oxidation reactions [26]. This distribution of active copper in equal proportions among ceruloplasmin and albumin only occurs in the postabsorptive period. In the postprandial period albumin is highly enriched in copper, as it functions as a short-lived metabolic compartment for adsorbed copper in its transport from the gastrointestinal tract to the liver [27,28]. In humans, a significant increase in plasma cystine without a concomitant increase in Cys concentration has been reported in the postprandial state [8]. In the postabsorptive state, exportation of amino acids from peripheral tissues is associated with the appearance of cystine but not of Cys [8]. These two observations indicate that Cys from dietary proteins or endogenous pro-
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tein catabolism rapidly oxidizes in plasma, in fact, Cys presents an exceptionally low plasma concentration in comparison with other protein-forming amino acids [29]. These physiological data are consistent with our results, and support the hypothesis that albumin plays a critical role in Cys oxidation through the redox cycling activity of its bound copper atom, mainly in the postprandial period when portal blood is enriched in Cys generated from protein digestion and in copper bound to albumin. The mechanism of Cys oxidation proposed in this work would help explain two aspects of albumin metabolism, which to date have been poorly defined. One aspect would be the formation of nonmercaptalbumin in the circulation, and the other the signaling pathway for albumin degradation. Although albumin is secreted by the liver with its thiol group fully reduced, circulating albumin has 60% of the thiol groups forming a disulfide bond with Cys (nonmercaptalbumin). According to our scheme, cystine generated during Cys oxidation could supply the oxidative equivalents driving oxidation of the albumin thiol group. In fact, this reaction based on thiol disulfide exchange has been used for years for preparing nonmercaptalbumin [30]. Copper catalyzed oxidation of Cys leads to the generation of several reactive oxygen species in a complex process with two-phase kinetics [22]. Modification of secondary and tertiary structures and increased proteolytic susceptibility have been described in albumin exposed to oxygen radicals [31,32]. These radicals could damage albumin in the surroundings of the copper-binding site due to their high reactivity, and the resulting modifications could constitute the signal for selecting an albumin molecule for degradation via binding to scavenger receptors and posterior uptake into endocytic vesicles [3]. Acknowledgment The author thanks Nuria Martin for technical assistance. References [1] J. Masuoka, J. Hegenauer, B.R. Van Dyke, P. Saltman, J. Biol. Chem. 268 (1993) 21533–21537. [2] A. Zgirski, E. Frieden, J. Inorg. Biochem. 39 (1990) 137–148. [3] T. Peters Jr., All about Albumin. Biochemistry, Genetics and Medical Applications, Academic Press, San Diego, 1996. [4] G.J. Quinlan, C. Coudray, A. Hubbard, J.M.C. Gutteridge, J. Pharm. Sci. 81 (1992) 611–614. [5] S. Sengupta, C. Wehbe, A.K. Majors, M.E. Ketterer, P.M. DiBello, D.W. Jacobsen, J. Biol. Chem 276 (2001) 46896–46904. [6] M.A. Mansoor, C. Bergmark, A.M. Svardal, P.E. Lønning, P.M. Ueland, Arterioscler. Thromb. Vasc. Biol. 15 (1995) 232–240. [7] P.M. Ueland, M.A. Mansoor, A.B. Guttormsen, F. Mu¨ller, P. Aukrust, H. Refsum, A.M. Svardal, J. Nutr. 126 (Suppl. 4) (1996) 1281S–1284S.
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