Heme-induced heme oxygenase-1 (HO-1) in human monocytes inhibits apoptosis despite caspase-3 up-regulation

Detlef Lang1, Stefan Reuter1, Tania Buzescu1, Christian August2 and Stefan Heidenreich1

1 Department of Medicine D and 2 Institute of Pathology, University of Muenster, Muenster, Germany

Correspondence to: D. Lang; E-mail: langd{at}uni-muenster.de


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Monocyte activation, apoptosis and differentiation are hallmarks of most inflammatory vascular disorders. We studied the effects of heme oxygenase-1 (HO-1) induced by its substrate hemin on apoptosis, caspase-3 expression and the differentiation of freshly isolated human monocytes. Hemin induced HO-1 in a dose- and time-dependent fashion as measured by semi-quantitative RT–PCR and flow cytometry. Apoptosis was markedly suppressed by hemin in cells rendered apoptotic by serum deprivation or dexamethasone as determined by flow cytometric detection of annexin V binding or transmission electron microscopy (TEM). The specific HO-1 inhibitor zinc protoporphyrin (ZnPP) reversed the effects of hemin on monocyte apoptosis and diminished cell lifespan. Surprisingly, the cytoprotective effects of hemin were positively correlated with caspase-3 up-regulation. Hemin-induced apoptosis suppression was enhanced by the caspase-3 inhibitor DEVD-CHO, indicating that caspase-3 was active in a pro-apoptotic fashion. Hemin inhibited CD95 as a putative cytoprotective mechanism. Morphological studies and detection of CD86 showed that monocytes differentiated into macrophages in response to hemin after relatively long incubation times, a phenomenon that might be provoked by caspase-3-regulated pathways. Our results confirm a similar cytoprotective effect of hemin/HO-1 for monocytes as has been shown for other cells, despite caspase-3 up-regulation. The fact that HO-1 may adversely affect monocyte survival and differentiation could be of particular significance in future therapies for occlusive vascular diseases or transplant rejection.

Keywords: apoptosis, caspase-3, heme oxygenase-1, monocytes


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Heme oxygenase-1 (HO-1) is one of three enzyme isoforms with the capacity to degrade tissue heme (1,2). Heme oxygenases (HO) are able to open the heme ring and convert heme into biliverdin, triggering the release of iron and carbon monoxide. This is the initial and rate-limiting step in heme degradation. HO-1 is the inducible HO isoform that responds rapidly to diverse stimuli (3) and protects tissues against a wide range of injuries including nephrotoxic (4), hypoxic (5), hyperoxic (6), ischemic (7) and inflammatory (8) disorders. These protective effects of HO-1 result from enzyme actions that block injury pathways by, for example, reducing heme expression, a pro-oxidant species (9) or by generating carbon monoxide, a gaseous product that is vasodilatory, anti-apoptotic and anti-inflammatory (6,10). In addition, biliverdin and bilirubin have antioxidant and cytoprotective effects that may enhance HO-1 effects (11).

The significance of HO-1 expression for protection against injury to and counter-regulation of apoptotic pathways has not been elucidated for freshly isolated human monocytes. Monocytes/macrophages play a central role in the host's innate and acquired immunity by providing protection against infectious micro-organisms and tumours. Monocytes in a culture environment can be rescued from constitutive occurring apoptosis through serum supplementation or the addition of growth factors (12,13). The survival or apoptosis of monocytic cells can be crucial factors in determining whether an inflammatory disease spreads or subsides. Gaining insight into the mechanisms involved in the induction or inhibition of apoptotic events could also have therapeutic implications. The present study investigated the role of hemin-induced HO-1 as a putative protection factor against monocyte apoptosis. We elucidated the involvement of caspase-3, since caspase-3 performs a number of executive functions leading to apoptosis.


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Antibodies and reagents
The following materials were obtained: phycoerythrin (PE)-conjugated mAb against activated caspase-3 (#557091) (Pharmingen, San Diego, CA); caspase-3 inhibitor (CPP 32, DEVD-CHO) (Calbiochem, Bad Soden, Germany); PE-conjugated mouse anti-human Leu M3 mAb (anti CD14, clone P9, IgG2b) and control mAb of appropriate Ig isotypes and antibodies against bcl-2 (clone bcl-2/100), HO-1 (clone HO-1-1), CD95 (clone DX2) and CD86 (clone 2331) (Becton Dickinson, Palo Alto, CA); FITC-labeled annexin V (Bender Medsystems, Vienna, Austria); FITC-labeled F(ab')2 fragments of goat anti-mouse IgG (Dianova, Hamburg, Germany); caspase-3 antibody (#9662) detecting inactive pro-caspase-3, for western blotting analysis (Cell Signaling, Beverly, MA); reverse transcriptase (Stratagene, Heidelberg, Germany); Taq DNA polymerase (Gibco BRL, Karlsruhe, Germany); dNTPs (New England Biolabs, Beverly, MA); pd(N)6 (Boehringer Mannheim, Mannheim, Germany); agarose (AGS, Heidelberg, Germany). Unless otherwise indicated, all other reagents, as well as hemin (# H-5533), were obtained from Sigma Chemical Co. (St Louis, MO).

Monocytes and cell culture
Human monocytes were isolated from the leukocyte buffy coats of healthy volunteers. Mononuclear cells were obtained by Ficoll-Hypaque density gradient centrifugation (400 g, 20 min) following which they were washed and then purified further by centrifugation on a hypotonic Percoll density gradient [57% in phosphate buffered saline (PBS); 400 g, 30 min]. Two interphases were detected whose upper phase contained enriched monocytes. Cells were harvested, washed three times in cold PBS and seeded out into 24-well culture dishes (Greiner, Nürtingen, Germany) in RPMI 1640 culture medium (CM) containing 2 mM L-glutamine, 50 µg/ml penicillin/streptomycin, 5 mM HEPES and 10 µM mercaptoethanol at 37°C in a 5% CO2/95% air atmosphere. Monocytes were further purified by adherence to the culture dishes, resulting in a final purity of >85%. This was assessed by flow cytometry on a FACScan flow cytometer (Becton Dickinson, Mountain View, CA) defined by forward and side light scatter properties, as well as by staining for nonspecific esterase and detection of the CD14 surface molecule. Monocytes (0.5 x 10/ml) were incubated in CM with 5% fetal calf serum (FCS; endotoxin content <0.01 ng/ml) or were rendered apoptotic by serum starvation with 0.2% FCS for the indicated culture time. The results were not affected by LPS contamination since all reagents showed <0.001 EU/ml endotoxin concentration on limulus amebocyte lysate assay (BioWhittaker, Walkersville, MD).

Detection of apoptosis
Flow cytometric quantification of apoptosis.
Monocyte apoptosis was detected either with propidium iodide (PI) nuclei staining or through the use of flow cytometry which detects CD14 expression and annexin V binding as described previously (12,14). Simultaneous detection of CD14 expression and annexin V binding identifies apoptotic monocytes. We have shown in previous studies that monocytes down-regulate CD14 expression before becoming apoptotic (14). Monocytes were deemed to be apoptotic cells when CD14 expression was low and annexin V binding was high after lymphocytes were gated out flow cytometrically by forward and side light scatter properties. Monocytes prepared and treated as described above were double labelled with PE-conjugated Leu M3 mAb and annexin V–FITC in staining buffer (SB containing 1% BSA in 50 mM HEPES buffer, pH 7.4) for 15 min on ice. PE- and FITC-conjugated murine IgG mAb of unrelated specifities were used as controls. Cells were washed after staining and fixed in 4% paraformaldehyde to apply flow cytometry for a total of 10 000 events. Apoptosis was confirmed by DNA electrophoresis and TEM.

Transmission electron microscopy (TEM).
Typical morphologic changes indicative of apoptosis were evaluated by electron microscopy as described previously (13). Cells were washed off the culture dishes, centrifuged, fixed in 1% glutaraldehyde/0.1 M Na cacodylate–HCl, pH 7.4 and postfixed in 1% OsO4/0.15 M Na cacodylate–HCl, pH 7.4. Samples were dehydrated in an ascending ethanol series and embedded in epoxy resin (Epon 812). Ultrathin sections were mounted on 150 mesh Formvar-coated copper grids, post-stained with aqueous saturated uranyl acetate and 2% lead citrate, and were then examined using a Philips CM 10 electron microscope (Philips Electronics, Mahway, NJ) at an accelerating voltage of 60 kV.

Detection of apoptosis using DNA electrophoresis.
DNA extraction and electrophoresis were performed as described previously (14). 1 x I06 monocytes were lysed by a hypotonic lysing buffer (10 mM Tris, 1 mM EDTA and 0.2% Triton X-100, pH 7.4). After centrifugation (13 000 g, 30 min), supernatants containing cleaved chromatin were treated with RNase (50 pg/ml) and proteinase K (100 pg/ml). DNA was then extracted using phenolichloroform. After precipitation was performed with –20°C ethanol and the samples were dried and heated, equal amounts of DNA were placed on a 1% agarose gel and separated by electrophoresis for 1 h at 80 V. The lower detection limit for visualization of oligonucleosomal bands was 1.0 pg of DNA.

Evaluation of cell necrosis
Monocyte viability following all numbers of treatments was determined by trypan blue exclusion or PI uptake of nonpermeabilized cells using flow cytometry.

Flow cytometric detection of HO-1, caspase-3, CD95, bcl-2 and CD86
Expression of the aforementioned receptors and proteins was measured by flow cytometry. Monocytes were washed in PBS, fixed with 4% paraformaldehyde and permeabilized with 0.1% saponin for the purposes of detecting caspase-3, bcl-2 and HO-1. For staining, mAbs against activated caspase-3 (PE-conjugated rabbit anti-human), HO-1 [mouse anti-human, unlabeled IgG1, secondary labeling with affinity pure F(ab')2 goat anti-mouse IgG], bcl-2 (FITC-labeled mouse anti-human), CD95 (PE-conjugated mouse anti-human) or CD86 (FITC-conjugated mouse anti-human) (5 µg/ml each) or isotype-matched control murine mAb were applied for 20 min.

Indirect immunofluorescence analysis of HO-1 expression in monocytes
Monocytes were plated on sterile coverslips (LAB-Tek II, Nunc, Wiesbaden) in 24-well plates at 2.5 x 105 cells/ml. After 60 h of culture, adherent monocytes were fixed in 500 µl methanol. Following a further three washes in PBS, the coverslips were incubated for 10 min in heat-inactivated FCS serum (1/10) to block non-specific antibody binding to Fc receptors. To visualize HO-1 in monocytes, cells were then incubated at room temperature 2 h with mAb specific for HO-1. The cells were washed and then labeled by incubating for 90 min with a Cy3 labeled secondary antibody. After a further three washes, cells were incubated with DAPI reagent, washed again and examined under oil immersion microscopy using a immunofluorescence microscope (BX 60, Olympus optical) with a 3-CCD camera (DXC-950P, Sony, Köln).

Western blotting analysis
The cells were washed in PBS and resuspended at 10 cells/100 µl of sample buffer containing 2% SDS, 62.5 mM Tris HCl (pH 6.8), 10% glycerol, 5% ß-mercaptoethanol and bromphenol blue. The cells were then heated at 95°C for 10 min and stored at –20°C until analysis. Each sample contained 20 µg protein. The samples were separated by a NuPAGE gel 4–12% BT (Invitrogen, Groningen, The Netherlands) and transferred to nitrocellulose membranes [BA83 (0.2 µm) Schleicher und Schüll, Dassel, Germany] using a western blotting module (Invitrogen). The membranes were washed three times with potassium phosphate buffer (0.05 M K3PO4, pH 8.5) and were then incubated with DIG-NHS-ester (digoxigenin-3-O-methylcarbonyl-{epsilon}-aminocapron acid-N-hydroxy-succinimidester) and Nonidet P 40 (0.01% v/v) for 2 h. After being blocked with casein blocking reagent (Bio-Rad) for 30 min, the membranes were incubated with anti-caspase-3 antibody (1:1000). Following incubation with anti rabbit IgG peroxidase for 2 h at room temperature and incubation with Pierce Supersignal Femto West, reactive bands were visualized for the purposes of detecting caspase-3. Mouse IgG2b was used as an isotype matched control for primary antibody. Visualized bands were analysed using a Roche Lumi-imager F1.

Inhibition of caspase-3
A specific caspase-3 inhibitor (CPP 32, DEVD-CHO) was used at a concentration of 100 nM for the purposes of inhibiting caspase-3. Cells were prepared as described above and incubated during the entire culture period.

Semiquantitative RT–PCR
Post-stimulation RNA isolation from monocytes was conducted using an RNeasy Kit (Quiagen, Hilden, Germany) in accordance with the manufacturer's instructions. DNase (DNase I, RNase-free, Boehringer Mannheim, Germany) digestion was performed prior to transcription to cDNA, which was synthesized following the addition of 5 µM random primers [pd(N)6, Roche Diagnostics, Mannheim, Germany], 1 mM dNTPs (NEB, Beverly, MA) and incubation at 37°C with moloney murine leukemia virus reverse transcriptase (Stratagene, Heidelberg, Germany). Contamination with DNA was excluded by performing PCR from templates incubated without reverse transcriptase. The primers used for PCR amplification were 5'-ATGGATGATGATATCGCCGCG-3' and 5'-TCTCCATGTCGTCCCAGTTG-3' (human ß-actin, 248 bp) (13), as well as 5'-AAGATTGCCCAGAAAGCCCTGGAC-3' and 5'-AACTGTCGCCACCAGAAAGCTGAG-3' (human HO-1, 399 bp) (15). The PCR reaction mixture (40 µl) contained 2 mM MgCl2, 0.2 mM dNTP, 1 µM primer and 1 U Taq DNA polymerase. Samples were amplified by means of 30 cycles of 60 s denaturation at 94°C, 30 s annealing at 60°C (HO-1) or 55°C (ß-actin) and 60 s elongation in a Peltier thermal cycler (Biometra Uno II Thermocycler, Biometra, Göttingen, Germany).

The correlation between the expression levels of ß-actin and HO-1 was analysed for semiquantitative PCR. Signal intensity as measured by PCR products was analysed on a 1.5% agarose gel and visualized by ethidium bromide staining. Densitometric quantification of PCR signals was performed using the Bio Image Intelligent Quantifier program (Bio Image, Ann Arbor, MI).

Statistical analysis
Results are given as means ± SEM. The Mann–Whitney U test and (for paired comparisons) Wilcoxon signed rank test or Student's t-test were applied. P < 0.05 was deemed to be statistically significant. All experiments were performed at least five times with different buffy coats.


    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Hemin-induced down-regulation of monocyte apoptosis
Monocyte apoptosis was highly suppressed by hemin stimulation (Fig. 1). Apoptosis was quantified by the detection of CD14 and annexin V. Cellular apoptosis was induced by serum starvation and detected after 60 h of cell culture. Under control conditions (5% FCS), 7 ± 3.6% of monocytes were apoptotic. Serum reduction to 0.2% FCS yielded 55.7 ± 14.5% apoptotic cells. A hemin concentration of 2.5 µM down-regulated monocytic apoptosis (10.9 ± 3.6% apoptotic monocytes, P < 0.05 compared with 0.2% FCS). LPS (1 µg/ml) was used as a control and reduced apoptotic monocytes to 2.8 ± 1.1%.



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Fig. 1. Down-regulation of monocyte apoptosis by hemin. The amount of apoptotic monocyte was measured by flow-cytometric staining of CD14 and annexin V. Under control conditions (5% FCS) 7.0 ± 3.6% of cells were apoptotic. Serum reduction (0.2% FCS) induced apoptosis rates of 55.7 ± 14.5%. This was significantly reduced by stimulation with hemin (10.9 ± 3.6% apoptosis) which yielded survival rates comparable with those for LPS as a monocyte activator suppressing apoptosis (2.8 ± 1.1% apoptosis). LPS contamination was excluded since polymyxin B did not revert the effects of hemin. These data are means ± SEM from 12 independent experiments (*P < 0.05).

 
Comparable findings were obtained from the evaluation of monocytes via DNA electrophoresis and TEM (Fig. 2). Hemin reduced DNA laddering following serum starvation (Fig. 2A). Figure 2(B) shows representative TEM images of monocytes under different stimulation conditions. Approximately two-thirds of the monocytes in each visual field were apoptotic following serum starvation. Hemim stimulation reduced apoptotic monocytes to 10–15% of cells per visual field (P < 0.05). Although only negligible numbers of cells showed morphological changes characteristic of apoptosis, some specific changes following hemin stimulation under serum starvation were readily apparent. The cells formed tight clumps with some focus on nucleus staining. However, this did not impede detection of surface molecules, since cells were smoothly detached from each other for FACS analysis. The cells showed the same CD14 expression levels as for the control cells (5% FCS). Thus, it is possible that morphological changes are not attributable to apoptosis and that they are instead elicited by unspecific stress-induced changes secondary to serum starvation and hemin stimulation. All experiments were performed at least five times.



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Fig. 2. Down-regulation of apoptosis after stimulation with hemin detected by DNA electrophoresis and TEM. (A) Oligonuclosomal DNA cleavage of monocytes after hemin (2.5 µM) treatment. Electrophoresis of DNA isolated from 1 x 106 monocytes was performed after 48 h culture. For medium control (5% FCS), no nucleosomal DNA fragments were detected after ethidium bromide staining. After serum starvation (0.2% FCS), typical oligonucleosomal DNA laddering was found. Oligonucleosomal DNA fragments were significantly reduced after hemin (2.5 µM) treatment. (B) TEM demonstrates protection from apoptosis by hemin. The left section depicts two monocytes that maintained normal morphology after a 60 h culture in 5% FCS containing medium. The middle panel shows typical changes of two apoptotic monocytes after serum deprivation for 60 h. Dense nuclear condensation, cytoplasmic vacuolization, shrinkage and rounding are visible in the upper monocyte. The lower monocyte shows an early stage of apoptosis with incipient nuclear condensation. Less than 15% of the monocytes analysed showed typical apoptotic features per visible field after serum deprivation and concurrent stimulation with hemin (2.5 µM; right panel). No typical changes of apoptosis are discernible, although some unspecific changes are remarkable. The cells clumped and had had some emphasis in nuclear staining. These were unspecific changes owing to stress reaction. Magnification is x7200.

 
Time- and dose-dependent induction of monocytic HO-1 by hemin at mRNA and protein levels
Figure 3(A and B) shows that HO-1 mRNA was induced by hemin in monocytes in a dose- (A) and time-dependent (B) manner. This was done by semiquantitative RT–PCR whereby the resulting means ± SEM of arbitrary units after densitometric quantification of PCR signals are shown in a graph of one representative RT–PCR blotting for five experiments (Fig. 3A). Signal intensity was compared to the ß-actin level analysed by semiquantitative RT–PCR. Hemin induced HO-1 significantly from the basal level of 1 arbitrary unit after 24 h of incubation time up to 2.7 ± 1.6 units (1 µM hemin; P < 0.05). HO-1 mRNA increased continuously with increased dosages of hemin and led to a maximum mRNA intensity of 9.4 ± 3.4 units with 10 µM hemin.





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Fig. 3. Dose and time dependency of HO-1 mRNA expression (A and B) and of HO-1 protein expression (C–G) after hemin treatment. HO-1 is induced by hemin in monocytes in a dose- (A) and time-dependent fashion (B) at the mRNA and protein level (C–G). Hemin at a concentration of 1 µM induced HO-1 significantly after 24 h of incubation time (P < 0.05). HO-1 mRNA increased continuously with increased dosages of hemin, resulting in a maximum of mRNA intensity of 9.4 ± 3.4 units with 10 µM hemin (A). HO-1 mRNA peaked at 24 h of incubation and decreased again after 48 h. A significant increase was measurable after 12 h, resulting in 4.7 ± 2.0 arbitrary units. This enhancement in expression nearly doubled after 24 h (7.9 ± 2.2 arbitrary units) (*P < 0.05) (B). HO-1 signal intensity was compared to ß-actin analysed by semiquantitative RT–PCR. One representative RT–PCR blotting is shown out of five experiments. The HO-1 protein level showed comparable regulation to mRNA (C–F). Hemin (1 µM) stimulated HO-1 protein expression significantly detected by FACS analysis. Time dependency was detected at protein level at different times than mRNA since protein expression has to be expected during a later time course. (C and D) Quantitation and statistical analysis of eight experiments are presented in (F) for different hemin concentrations and in (E) for time dependancy (n = 5). For each of them one representative FACS analysis is shown in (D) and (F), respectively. HO-1 protein expression peaked at 16 h of stimulation with hemin (*P < 0.05). Induction of HO-1 was also detected by indirect immunofluorescence after 60 h (G). Without stimulation nearly no HO-1 expression was detectable (red signals, upper panel). Stimulation with 1 µM hemin (middle) and 2.5 µM hemin (lower panel) resulted in increasing signals for HO-1 expression and confirmed FACS Data. Iso = isotype control.

 
Since 2.5 µM hemin increased HO-1 mRNA significantly, this concentration was used to investigate time dependency. HO-1 mRNA peaked at 24 h of incubation and declined after 48 h. After 2 h, only a moderate increase in HO-1 mRNA (1.3 ± 0.1 arbitrary units) was detected. A significant increase was observed after 12 h, resulting in 4.7 ± 2.0 arbitrary units, and this increase nearly doubled after 24 h (7.9 ± 2.2 arbitrary units) (*P < 0.05; Fig. 3B).

HO-1 regulation at protein level was comparable to mRNA expression (Fig. 3C and D). One-micromolar hemin stimulated HO-1 protein expression significantly from a basal level of 11.9 ± 5.7% cells expressing HO-1 up to 40.2 ± 13.3%, as quantified by FACS analysis (P < 0.05). Time dependency was detected at protein level at different times than mRNA, since protein expression has to be expected during a later time course (E and F). Flow-cytometric analysis revealed that HO-1 protein levels increased continuously with increased dosages of hemin and peaked after 16 h of hemin stimulation (2.5 µM) (70 ± 9% HO-1 positive cells) (*P < 0.05; Fig. 3C and E).

Hemin-induced caspase-3 and the effects on apoptosis of additional caspase-3 inhibition
Our analysis of caspase-3 expression as one of the main effectors of apoptosis revealed that hemin significantly induced activated caspase-3. Culturing cells in 0.2% FCS containing medium caspase-3 positive cells increased from 23 ± 4% (0.2% FCS) to 56.6 ± 6 % (2.5 µM, P < 0.05) (Fig. 4A). In order to rule out the possibility that the increase in the amount of activated caspase-3 positive cells was elicited by increased precursor induction, we performed western blotting analysis against pro-caspase-3 (Fig. 4C). Pro-caspase-3 was down-regulated following hemin stimulation, thus demonstrating that the increased caspase-3 level was mainly caused by activated/active caspase-3. The fact that caspase-3 was blocked by the specific inhibitor DEVD-CHO showed that hemin-induced suppression of the apoptosis rate could be further down-regulated (P < 0.05; Fig. 4D). Caspase-3 inhibition alone did not affect apoptosis induction following serum starvation with the concentration applied (56 ± 15% apoptosis without DEVD-CHO as compared to 51 ± 18% with DEVD-CHO). However, this concentration was combined with a hemin treatment that has the capacity to reduce apoptosis.



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Fig. 4. Induction of activated caspase-3 in monocytes by hemin (A and B) was paralleled by down-regulation of inactive pro-caspase-3 (C) when caspase-3 antagonist DEVD-CHO influenced hemin-dependent monocyte apoptosis (D). (A and B) Hemin significantly induced activated caspase-3. At 0.2% FCS containing medium, 23 ± 4% caspase-3 positive monocytes increased to 56.7 ± 6% by hemin stimulation (2.5 µM, P < 0.05). Monocytes cultured in 5% FCS served as control. A maximum increase was noted for 5 µM hemin. (B) Representative FACS data are shown for which quantification and statistical analyses are presented in (A). (C) Proving results generated by FACS analysis, western blotting in order to detect pro-caspase-3 with a specific antibody against inactive, uncleaved pro-caspase-3 was performed. Hemin down-regulated pro-caspase-3 after serum starvation (0.2% FCS and hemin) which demonstrated that FACS analysis detected activated caspase-3 and not inactive pro-caspase. (D) Blocking of active caspase-3 by the specific inhibitor DEVD-CHO (100 nM) showed that inhibition of apoptosis after hemin treatment could be further down-regulated (*P < 0.05, B). The relevant level of active caspase-3 was detected after 16 h, apoptosis after 60 h.

 
Effect of the HO-1 antagonist zinc protoporphyrine (ZnPP) on apoptosis
HO-1 inhibition by ZnPP, a competitive blocker of HO-1, led to a 30 ± 9% increase in the apoptosis rate in monocytes treated with hemin (P < 0.05) whereas ZnPP reversed the anti-apoptotic effects of HO-1, with 10.9 ± 3.6% of the cells becoming apoptotic following serum deprivation and HO-1 induction via hemin alone (Fig. 5). Dexamethasone-induced apoptosis showed apoptosis rates of 83.5 ± 2% that were not significantly altered by ZnPP. Hemin significantly reduced apoptosis due to the addition of dexamethasone (25 ± 5.7% apoptosis) and ZnPP again reversed this effect, leading to high levels of apoptosis (78.5 ± 3.5% apoptosis, P < 0.05). These results once again show that ZnPP can block hemin-induced anti-apoptotic pathways, and this in turn points to the cytoprotective effects of HO-1.



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Fig. 5. Influence of ZnPP as an HO-1 blocker on monocyte apoptosis induced by serum starvation alone or with dexamethasone treatment. ZnPP as a competitive blocker of HO-1 reversed the anti-apoptotic effects of HO-1 induced by hemin. Apoptosis increased significantly from 7.0 ± 3.0% to 32 ± 10% by ZnPP (10 µM) in 5% FCS-containing medium. The amount of apoptosis was not affected by ZnPP after serum starvation. A significant reduction of apoptosis was found in monocytes after hemin treatment, an effect which was significantly reverted again by ZnPP. Very high apoptosis levels achieved by 0.2% FCS medium plus dexamethasone could be highly suppressed by hemin and also reverted again by ZnPP. Dexa: dexamethasone at a concentration of 10 µM. The hemin concentration used was 2.5 µM. ZnPP: zinc protoporphyrine (10 µM). (*P < 0.05 compared to stimulation without ZnPP, + P < 0.05 compared to 0.2% FCS alone.)

 
Detection of CD95, bcl-2 and CD86
The primary CD95 death receptor is significantly down-regulated by hemin (22 ± 4% CD95 positive monocytes without hemin versus 10 ± 3% CD95 positive cells with hemin; P < 0.05) (Table 1). No significant differences were found for bcl-2, which is an important member of a family of proteins that prevents apoptosis. On the other hand, the costimulatory receptor CD86 significantly increased following hemin stimulation (4 ± 4% CD86 positive cells without hemin versus 19 ± 3% CD86 positive cells with hemin; P < 0.05), thus indicating that hemin is associated with the induction of monocyte differentiation into macrophages. Furthermore, we could show that the inhibition of HO-1 by ZnPP reverted the induction of CD86 by hemin. The same was true for inhibition of caspase-3 which also inhibited the expression of hemin-induced costimulatory CD86 (Fig. 6).


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Table 1. Regulation of CD95, bcl-2 and CD86 by hemin treatment

 


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Fig. 6. Induction of CD86 in monocytes by hemin was abrogated by ZnPP as an HO-1 blocker and the specific caspase-3-inhibitor DEVD-CHO. A marker for differentiation in human monocytes is CD 86. Hemin could induce significantly the expression of CD 86 after 168 h of incubation whereas ZnPP (10 µM) as an HO-1 blocker and the specific inhibitor Caspase-3-Inhibitor (C-3-Inhib) DEVD-CHO (100 nM) themselves did not change expression of CD 86 compared to unstimulated monocytes (ctrl = control). ZnPP, as well as the caspase-3-inhibitor, could also prevent the expression of CD86 after stimulation by hemin. Influence of relevant endotoxin contamination was excluded by additional polymyxin D stimulation. (A) Four experiments are summarized. (B) A representative histogram.

 

    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Monocytes/macrophages are centrally involved in the pathogenesis of various inflammatory and vascular disorders such as sepsis, atherosclerosis and transplant rejection (1618). Recent data from organ transplantation, xenotransplantation, ischemia/reperfusion injury and inflammatory kidney disorders reveals that the HO-1 enzyme diminishes inflammation and improves outcome in these disorders by mediating cytoprotection in various cell types (1921). Most studies have investigated the effects of HO-1 on cardiac myocytes, endothelial cells or smooth muscle cells (2224). Few data are available on monocytes (25). HO-1 has become a focal point in a number of animal studies because it is easily inducible by its own substrate hemin/heme. HO-1 can also be blocked by metallo-protoporphyrins, all of which have minor toxic effects when pharmacological doses are applied in vivo (2629). In view of the crucial role played by monocytes, particularly in the aforementioned settings, we elucidated the effects of HO-1 in freshly cultured cells on the mechanisms and regulation of apoptosis.

As was found for other cell types (2), hemin is a highly effective HO-1 inducer that antagonized PCD effectively in monocytes rendered apoptotic by serum deprivation or dexamethasone (Figs 1 and 5). We found that this cytoprotective effect is reversed by the specific competitive HO-1 inhibitor, ZnPP. This indicates that hemin-enhanced life span is associated with induction of active HO-1, establishing a direct link between HO-1 up-regulation and cytoprotective effects.

Hemin up-regulated monocyte HO-1 dose-dependently on the mRNA and protein levels. HO-1 peaks were obtained after 16 and 24 h, but apoptosis levels could only be measured after 60 h owing to the fact that the cells did not induce apoptotic changes before that time. We found that the anti-apoptotic effect of hemin treatment was most pronounced when hemin was added to the culture medium within 24 h. Hemin added at a later time was less protective (data not shown). We also tried to ascertain which HO-1-dependent metabolic products (iron, CO and bilirubin or biliverdin) trigger cytoprotection in monocytes. Since iron sulfate/ferric gluconate and the iron chelator desferrioxamine had no effect on hemin-induced protection from PCD, iron-related effects could be ruled out. CO and bilirubin were not measurable in our culture supernatants or cell lysates due to the extremely low concentrations or short half-life times of these substances. Thus, we were unable to elucidate the role of these two metabolites. However, exogenously administered bilirubin significantly inhibited apoptosis of monocytes (data not shown), which suggests that this substance may mediate hemin-dependent cytoprotection. Other key mechanisms such as reactive oxygen intermediate pathways may also play a role in the regulation of spontaneous monocyte apoptosis blocked by HO-1. These pathways require further investigation. Surprisingly, our additional investigations of hemin-induced changes revealed that, despite apoptosis inhibition, caspase-3 expression was up-regulated in monocytes. Caspase-3 activation by HO-1 has recently been demonstrated for vascular smooth muscle cells, but here, adenovirus-mediated HO-1 stimulation triggered apoptosis instead of abrogating it (30). Hemin activated caspase-3 effectively in monocytes, which was proven by co-administration of the specific caspase-3 antagonist DEVD-CHO which further enhanced cell protection and the life span of monocytes in our investigation. In our view, the effects of hemin on apoptosis are regulated in a cell-specific manner and HO-1 may not have a protective effect in all types of cells.

Our findings demonstrate that the effects of hemin are not limited to HO-1 induction and caspase-3 activation; in our study, apoptosis-mediating receptors such as CD95 were also down-regulated (Table 1). This inhibition could be a principal cytoprotective effector in settings involving the CD95/CD95L system, as has been shown for glucocorticoid-induced monocyte PCD (31). Since hemin effectively counter-regulates apoptosis following serum starvation—a process that does not involve CD95/CD95L to any significant degree—other receptors and pathways must be involved. In our supplementary morphological studies of hemin-treated monocytes cultured for more than 60 h, rapid differentiation into macrophages was observed (Table 1), nuclei shape altered, cytoplasmic volume increased (Fig. 2) and expression of the costimulatory receptor CD86 (which is characteristic of macrophage activation) rose significantly (Table 1). Classic dendritic cell markers such as CD1a remained unaffected. This monocyte to macrophage differentiation might be attributable to hemin-induced caspase-3 up-regulation and HO-1 induction since specific inhibition of caspase-3 and HO-1, respectively, blocked the induction of CD 86 (Fig. 6).

A recent report revealed that the principal monocyte growth factor M-CSF induces differentiation of monocytes via caspase activation and apoptosis suppression involving cytochrome c release from mitochondria (32). In contrast to M-CSF-induced differentiation, hemin-dependent differentiation might be promoted by iron, which is released by HO-1 activation and has been attributed to monocyte into macrophage differentiation (33,34). Our findings are of particular significance in light of recent reports on the beneficial effects of HO-1-dependent anti-inflammation or tissue protection (19,35,36). In settings where vascular occlusive disorders or organ transplant rejection are amenable to amelioration by HO-1 up-regulation via smooth muscle cell apoptosis with concurrent consecutive vascular stenosis desobliteration and endothelial cell protection that promotes re-endothelialization, this improvement could be antagonized by HO-1-specific effects on monocytes. On the other hand, recent data show that the influx of monocytes/macrophages are not necessarily synonymous with tissue damage and progressive inflammation (37). Macrophage function may be more important than the number of infiltrating cells. Additional studies need to elucidate the changes in monocyte function by different treatments which will also help to understand the regulation of local inflammatory responses.

Nevertheless, in atherosclerosis, monocytes are supposed to aggravate lesions and promote stenosis and vascular remodeling. In organ transplantation, these cells seem to provoke immunosuppressive-resistant rejection or fibrotic injury (28,38). Supporting monocyte survival and differentiation into macrophages, hemin/HO-1 treatment may have more than just beneficial effects achieved by effects on intrinsic vascular cells during these processes. Thus, further studies on HO-1-dependent tissue protection should avoid inducing anti-apoptotic effects in monocytes whereby especially the function of monocytes has to be taken into account, probably more than the absolute number of infiltrating monocytes.

In conclusion, our study shows that hemin can generate HO-1 in cultured monocytes. Although HO-1 induction antagonizes apoptosis, there is an increased caspase-3 activity and a distinct induction of cell differentiation into macrophages. Experiments that treat various inflammatory, vascular or immune diseases with HO-1 should precisely characterize the effects of this enzyme on all cellular compartments involved.


    Acknowledgements
 
The authors thank Ms. K. Beul for excellent technical assistance. This work was supported by I2KF Muenster, Project No. D20.


    Abbreviations
 
CM   culture medium
FCS   fetal calf serum
HO   heme oxygenase(s)
HO-1   heme oxygenase-1
PCD   programmed cell death
PI   propidium iodide
PS   phosphatidylserine
SB   staining buffer

    Notes
 
Transmitting editor: T. Hunig

Received 23 March 2004, accepted 11 November 2004.


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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