mitochondrial transmembrane potential is diminished in ... transmembrane potential.pdfmitochondrial...

Original Paper

Mitochondrial transmembrane potential is diminished in phorbol myristate acetate-stimulated peritoneal resident macrophages isolated from wild-type mice, but not in those from gp91-phox-deficient mice Toshihiro Kobayashi1 , Yasuhiro Ogawa2, Yoshiya Watanabe3, Masato Furuya3, Sayo Kataoka4, Eva Garcia del Saz1, Shohko Tsunawaki5, Mary C. Dinauer6 and Harumichi Seguchi1

Accepted: 9 June 2004 Published online: 9 July 2004

Abstract Macrophages produce superoxide (O2–) during phagocytosis or upon stimulation with

a variety of agents including phorbol myristate acetate (PMA) through the activation of NADPH oxidase, and the formed O2

– is converted to other reactive oxygen species (ROS) such as hydrogen peroxide (H2O2). The aim of the present study was to elucidate the effect of the intracellularly produced ROS on mitochondrial transmembrane potential (MTP) in mouse (C57BL/6) peritoneal resident macrophages stimulated with PMA. Using a fluorescent dye, succinimidyl ester of dichlorodihydrofluorescein (H2DCFDA), O2

– was visualized in intracellular compartments in a certain subpopulation of macrophages isolated from wild-type

Histochemistry and Cell Biology © Springer-Verlag 200410.1007/s00418-004-0674-0

(1) Department of Anatomy and Cell Biology, Kochi Medical School, Kochi University, Kohasu, Okoh-cho, Nankoku, 783-5305 Kochi, Japan

(2) Department of Radiology, Kochi Medical School, Kochi University, Kochi, Japan(3) Institute for Laboratory Animals, Kochi Medical School, Kochi University, Kochi, Japan(4) Medical Research Center, Kochi Medical School, Kochi University, Kochi, Japan(5) Department of Infectious Diseases, National Research Institute for Child Health and Development,

Tokyo, Japan(6) Department of Pediatrics, Indiana University School of Medicine, Indianapolis, USA

Toshihiro Kobayashi Email: [email protected] Fax: +81-888-802304

of 2310.1007/s00418-004-0674-0

7/15/2004http://springerlink.metapress.com/media/LP3WUPQYMKP1BTM3NG86/Contributions/N...

mice. Cells deficient in gp91-phox, one of the membrane components of NADPH oxidase, were negative for the fluorescence. When cells were loaded with both H2DCFDA and MitoCapture, a

fluorescent dye for mitochondria, mitochondrial fluorescence was diminished in O2–-producing

cells, but not in O2–-deficient cells. Flow cytometry also revealed the decrease of mitochondrial

fluorescence in wild-type cells, but not in gp91-phox-deficient cells. The loss of mitochondrial fluorescence was prevented by microinjection of catalase into cells. The present findings demonstrate that MTP is diminished by ROS, including the H2O2 dismutated from O2

–, produced intracellularly by activation of the NADPH oxidase in mouse peritoneal resident macrophages stimulated with PMA.

Keywords Apoptosis - Macrophages - Mitochondria - NADPH oxidase - Superoxide

Introduction Superoxide (O2

–) is produced by NADPH oxidase activity in phagocytes such as neutrophils and macrophages. This enzyme comprises four cytosolic (Rac, p40-phox, p47-phox, and p67-phox) and three membrane (p22-phox, gp91-phox, and Rap1A) components. During phagocytosis, or upon stimulation with a variety of agents including phorbol myristate acetate (PMA), the cytosolic phox proteins translocate and associate with the membrane components leading to the activation of NADPH oxidase. The formed O2

– is converted into other reactive oxygen species

(ROS), such as hydrogen peroxide (H2O2), hydroxyl radical (·OH), singlet oxygen (1O2), and hypochlorous acid (HOCl), which play crucial roles in the host defense by microbial killing, and also cause injury to surrounding tissues (for reviews see Karnovsky 1994; Robinson and Badwey 1995; Babior 1999, 2004; Kobayashi and Seguchi 1999, 2001; Kobayashi et al. 2001; Seguchi and Kobayashi 2002). Protein p29 is additionally associated with NADPH oxidase in a manner that this component inactivates the oxidant, or alters signaling pathways affected by the oxidant, protecting NADPH oxidase from auto-oxidation (Leavey et al. 2002).

The mode of action of ROS involves direct interaction with specific receptors, and/or redox-activation of members of signaling pathways such as protein kinases, protein phosphatases, and transcription factors. In addition, ROS act in concert with intracellular calcium ions in signaling pathways regulating the balance of cell proliferation versus cell cycle arrest and cell death (Sauer et al. 2001). ROS are also known to activate mitogen/extracellular signal-regulated kinase (MEK)1/2 and extracellular signal-regulated kinase (ERK)1/2 (Xiao et al. 2002).

Morphological studies using human neutrophils have demonstrated that, upon cell stimulation, initial O2

– production occurs in special intracellular compartments. These oxidant-producing organelles then bind directly to the plasma membrane or fuse to form larger structures that eventually become associated with the plasma membrane, and O2

– is released extracellularly from the cells (Kobayashi and Robinson 1991; Kobayashi et al. 1998).

of 2310.1007/s00418-004-0674-0


Disruption of the mitochondrial transmembrane potential (MTP) followed by the subsequent release of cytochrome c has been suggested to be a critical mediator of apoptosis (Wadia et al. 1998; Hancock et al. 2001; Chauhan et al. 2003). In this regard, we have previously confirmed that ROS formation results in the decrease of MTP and in the release of cytochrome c from mitochondria in human peripheral blood T lymphocytes irradiated with X-rays (Ogawa et al. 2002, 2003a, b). ROS also reduce MTP, induce the release of cytochrome c, and activate caspase-9 and caspase-3 in a variety of cancer cells (Chung et al. 2003). Moreover, the activation of NADPH oxidase has been reported to induce apoptosis, as evidenced by apoptosis-specific phosphatidylserine externalization in human neutrophils (Fadeel et al. 1998) and in HL-60 cells (Arroyo et al. 2002).

To evaluate the localization of the oxidant-producing intracellular compartments we employed a cytochemical method previously developed in our laboratory (Kobayashi et al. 2000) using succinimidyl ester of dichlorodihydrofluorescein diacetate (H2DCFDA), which is not fluorescent until oxidized by ROS (Robinson et al. 1988). We further studied the disruption of MTP in T lymphocytes exposed to exogenously added H2O2 (Ogawa et al. 2004) using MitoCapture, a fluorescent dye, which is known to accumulate and aggregate in the mitochondria, giving off a red fluorescence in healthy cells, but not in apoptotic cells, and detects the disruption of MTP, one of the earliest intracellular events that occur following induction of apoptosis (Zamzami et al. 1995). The effect of ROS produced intracellularly by the activation of NADPH oxidase on MTP, however, remained to be established.

In the present study, cytochemical and biochemical approaches using H2DCFDA in combination with MitoCapture were employed to elucidate the effect of ROS on MTP, and it was demonstrated that MTP is diminished by ROS dismutated from O2

– produced intracellularly by the activation of NADPH oxidase in PMA-stimulated living macrophages obtained from mouse (C57BL/6) peritoneal cavity.

Materials and methods

Reagents

Phorbol 12-myristate 13-acetate (PMA) and catalase were purchased from Sigma (St. Louis, MO, USA). Succinimidyl ester of dichlorodihydrofluorescein diacetate (H2DCFDA), LysoTracker Blue, LysoTracker Red, and Cascade Blue-Dextran (MW 3000) were obtained from Molecular Probes (Eugene, OR, USA). MitoCapture was purchased from BioVision (Palo Alto, CA, USA). All other reagents were of the highest purity grade available.

Isolation of macrophages

Resident macrophages were isolated from peritoneal cavities of wild-type mice (C57BL/6) and gp91-phox-deficient knockout mice by the method of Badwey et al. (1983). Briefly, mice were killed by cervical dislocation under anesthesia with diethyl ether, and cell suspension was

of 2310.1007/s00418-004-0674-0


obtained from the peritoneal cavity after washing with ice-cold phosphate-buffered saline containing 1 mM MgCl2, 1 mM CaCl2, 5 mM glucose, and 0.2% bovine serum albumin. This buffer solution was also employed as washing and incubation solution throughout the experiment. Centrifuged cell pellets were resuspended in the buffer solution. Cells were attached onto a glass coverslip at the bottom of a microwell dish (35-mm dish, poly-D-lysine-coated; MatTek, Ashland, MA, USA) and used for the experiments described below after washing in the buffer solution (Kobayashi et al. 2000).

Cell stimulation

Macrophages attached onto a glass coverslip, immersed in 2 ml of the incubation buffer, were stimulated by treatment with PMA, a substance known to activate protein kinase C (Nishizuka 1986). Stock solutions of PMA were prepared in dimethyl sulfoxide (DMSO) and stored at –20°C. Cells were exposed to 1 M PMA for 30 min at 37°C. PMA stock solutions were diluted with

DMSO so that the final concentration of the solvent in the cell suspension was 0.25% (v/v). Unstimulated cells, which served as controls, were incubated in a similar fashion, but without the addition of PMA.

Treatment with fluorescent dyes

Cells were loaded for 30 min at 37°C with PMA and H2DCFDA at the final concentration of 25

M for the visualization of the O2–-producing site. MitoCapture was employed for the detection

of mitochondria according to the protocol commercially available. Cells exposed to MitoCapture were incubated for 30 min at 37°C with or without PMA. For the detection of lysosomal localization, cells were incubated for 30 min at 37°C in a medium containing PMA or without it, and LysoTracker at the final concentration of 50 nM was added to the medium 5 min before the completion of the incubation.

Microinjection of catalase into cells

Microinjections were performed using an Eppendorf Transjector 5246 and Eppendorf Micromanipulator 5171 (Hamburg, Germany) by the method of Fratti et al. (2001). Catalase was dissolved in 135 mM potassium chloride aqueous solution, and mixed with 2 mg/ml Cascade Blue-Dextran (MW 3000) to give a final catalase concentration of 20 mg/ml. Microinjections were carried out at a pressure of 50 hPa for 0.2 s using Eppendorf Femtotips II. The potassium chloride solution without catalase was employed as control.

Fluorescence microscopy

The microwell dish with attached cells immersed in 2 ml incubation buffer was placed on the thermoregulating stage of a fluorescence microscope (Axiovert S100TV; Zeiss, Jena, Germany) providing a constant temperature of 37°C. The microscope was equipped with a cooled CCD camera C4880 (Hamamatsu Photonics, Japan), a 100-W mercury lamp, and an appropriate set of filters. The exposure time of the CCD camera for detection of fluorescence was 0.2 s.

of 2310.1007/s00418-004-0674-0


Fluorescence-positive organelles were visualized using Aquacosmos software (Hamamatsu Photonics), and the image was composed with Adobe Photoshop (Adobe Systems, San Jose, CA, USA).

Cytochemical procedure for detection of NADPH oxidase activity

Oxidant-producing sites were visualized using a modification of the cerium-based cytochemical method (Kobayashi et al. 1999) of Briggs et al. (1975). Cells were washed with HEPES buffer (pH 7.4) consisting of 20 mM HEPES, 135 mM NaCl, 5 mM KCl, and 5 mM glucose, and incubated for 30 min at 37°C in HEPES buffer containing 20 mM tricine, 1 mM CeCl3, and 1 mM NaN3. Cells were then washed in HEPES buffer containing 20 mM tricine, and fixed with 2% glutaraldehyde in HEPES buffer for 10 min at room temperature.

Electron microscopy

After the cytochemical incubation, cells were postfixed for 10 min on ice with 2% osmium tetroxide and 1.5% potassium ferrocyanide in 0.1 M cacodylate buffer (pH 7.4) followed by dehydration through a graded series of ethanols, and embedding in Spurr s epoxy resin (Spurr 1969). Ultrathin sections (60 nm in thickness) were obtained in an Ultracut OmU4 ultramicrotome (Reichert-Jung, Vienna, Austria) and picked up on copper grids. These sections were poststained with lead citrate and uranyl acetate and observed under a JEM-100S (Jeol, Tokyo, Japan) operated at an accelerating voltage of 80 kV.

Flow cytometry

Quantification of mitochondrial fluorescence using MitoCapture was done according to the assay protocol commercially available. Briefly, cells (1×106/ml) were incubated in the incubation buffer containing PMA and MitoCapture for 30 min at 37°C, and then immediately assayed with a flow cytometer (FACScan; Becton Dickinson, San Jose, CA, USA). The fluorescent signal was detected using a channel for red fluorescence (Ex/Em=488/590 nm).

Results

Detection of O2–-producing intracellular compartments

We visualized O2–-producing sites in mouse peritoneal resident macrophages stimulated with

PMA using a fluorescent dye, H2DCFDA. Fluorescence was observed at intracellular compartments in PMA-stimulated cells (Fig. 1B), but not in unstimulated cells obtained from wild-type mice (Fig. 1A). The fluorescence was, however, restricted to a subpopulation of cells comprising approximately 20% of the total number of macrophages, as estimated by cell count

of 2310.1007/s00418-004-0674-0


under a fluorescence microscope, indicating that the macrophages are subdivided into two types: O2

–-producing and O2–-deficient cells. Fluorescence was not found in PMA-stimulated cells

isolated from knockout mice deficient in gp91-phox, one of the membrane components of NADPH oxidase, which consequently lack the ability to build up the active form of NADPH oxidase (Fig. 1C).

of 2310.1007/s00418-004-0674-0


of 2310.1007/s00418-004-0674-0


Fig. 1A–C Fluorescence micrographs of living peritoneal resident macrophages isolated from wild-type mice (A, B) and from gp91-phox-deficent mice (C). A Unstimulated cells exposed to succinimidyl ester of dichlorodihydrofluorescein (H2DCFDA). No specific fluorescence labeling was observed in these cells. B Phorbol myristate acetate (PMA)-stimulated cells exposed to H2DCFDA. Fluorescence was detected in intracellular compartments in a certain subpopulation of macrophages (closed arrows), but not in the other cells (open arrows). C PMA-stimulated cells exposed to H2DCFDA. No fluorescence was observed in the gp91-phox-deficient cells

Visualization of organelles by fluorescence microscopy

Different kinds of organelles including O2–-producing intracellular compartments, mitochondria,

and lysosomes were visualized in living macrophages, isolated from wild-type mice, under a fluorescence microscope using organelle-specific fluorescent dyes: H2DCFDA for O2

–-producing intracellular compartments, MitoCapture for mitochondria, and LysoTracker for lysosomes. O2

–-producing intracellular compartments (Fig. 2A) and lysosomes (Fig. 2C) were spherical in shape and of variable sizes. Mitochondria appeared as rod-shaped structures (Fig. 2B). This assay enabled us to visualize simultaneously the intracellular dynamics of plural organelles in living cells under a fluorescence microscope as reported below.

of 2310.1007/s00418-004-0674-0


of 2310.1007/s00418-004-0674-0


Fig. 2A–C Fluorescence micrographs of living peritoneal resident macrophages isolated from wild-type mice. A A PMA-stimulated cell showing fluorescence at superoxide (O2

–)-producing sites. Cells were exposed to H2DCFDA. B An unstimulated cell showing the localization of mitochondria. Cells were exposed to MitoCapture. C An unstimulated cell showing the localization of lysosomes. Cells were exposed to LysoTracker Blue. The fluorescence images were superimposed on the corresponding morphological micrographs. O2

–-producing intracellular compartments (A) and lysosomes (C) exhibit spherical shapes and are of different sizes. Mitochondria (B) appear as rod-shaped structures

Simultaneous detection of O2–-producing intracellular

compartments and mitochondria or lysosomes

The localization of O2–-producing intracellular compartments was concurrently investigated with

that of mitochondria or lysosomes in PMA-stimulated cells isolated from wild-type mice. Decrease in mitochondrial fluorescence was observed in O2

–-producing cells exposed to

H2DCFDA and MitoCapture. No fluorescence loss was, however, detected in O2–-deficient cells

(Fig. 3A–C): these cells showed identical intensity of mitochondrial fluorescence to that observed in unstimulated cells. When cells were treated with H2DCFDA and LysoTracker, O2

–-

producing cells exhibited lysosomal fluorescence similar to that of O2–-deficient cells (Fig. 3D–

F).

of 2310.1007/s00418-004-0674-0


of 2310.1007/s00418-004-0674-0


Fig. 3A–F Fluorescence and morphological micrographs of living peritoneal resident macrophages stimulated with PMA, isolated from wild-type mice. Cells were exposed to MitoCapture (A–C) for mitochondrial staining or to LysoTracker Red (D–F) for lysosomal staining, concomitantly with H2DCFDA. The open arrow in C indicates a cell exhibiting positive labeling for O2

– (A) and low mitochondrial fluorescence (B) as compared to the fluorescence in O2

–-deficient cells indicated by the closed arrows in C. The open arrow in F indicates a cell producing O2

– (D) with normal lysosomal fluorescence (E) similar to that observed in the O2

–-deficient cell indicated by the closed arrow in F

Flow cytometry for mitochondrial fluorescence Mitochondrial fluorescence was analyzed using a flow cytometer in cells isolated from wild-type and gp91-phox-deficient mice exposed to PMA and MitoCapture. The flow cytometric assay exhibited two peaks (100×2 FL2-H and 102×1 FL2H) with respect to fluorescence intensity of mitochondria in wild-type cells. The cell population of the minor peak (100×2 FL2-H) was estimated to represent 21% of the total cell number, and that of the major peak (102×1 FL2H) as comprising 79% (Fig. 4A). In gp91-phox-deficient cells, however, this minor peak (100×2 FL2-H) was not identified (Fig. 4B).

Fig. 4A, B Flow cytometric analysis of mitochondrial fluorescence in cells exposed to PMA and MitoCapture. A The intensity of mitochondrial fluorescence indicated the presence of two subpopulations of macrophages in wild-type mice (100×2 FL2-H and 102×1 FL2H). B In gp91-phox-deficient mice, however, the peak indicated with the open arrow in A was not identified

of 2310.1007/s00418-004-0674-0


Microinjection of catalase into cells

As shown in Fig. 3, mitochondrial fluorescence was diminished in O2–-producing, but not in O2

–

-deficient cells isolated from wild-type mice stimulated with PMA. It was, thus, considered that the MTP is disrupted by the O2

– produced intracellularly by NADPH oxidase activity. To confirm this, cells were microinjected with catalase which converts the H2O2 spontaneously

dismutated from O2– into O2 and H2O. The mitochondrial fluorescence in O2

–-producing cells microinjected with catalase was preserved to the same degree (Fig. 5A–C) as that observed in O2

–-deficient cells stimulated with PMA. As control, cells were microinjected with potassium chloride aqueous solution without catalase. As a result, the mitochondrial fluorescence was lost in O2

–-producing cells (Fig. 5D–F).

of 2310.1007/s00418-004-0674-0


of 2310.1007/s00418-004-0674-0


Fig. 5A–F Fluorescence and morphological micrographs of living peritoneal resident macrophages stimulated with PMA isolated from wild-type mice. Cells were microinjected with (A–C) or without catalase (D–F), and exposed to both H2DCFDA and MitoCapture. The closed arrow in C indicates a cell microinjected with catalase exhibiting O2

– production (A) and unchanged mitochondrial fluorescence (B). The closed arrow in F indicates a cell microinjected without catalase exhibiting O2

– production (D) and diminished mitochondrial fluorescence (E)

Electron microscopy The oxidant-producing sites and the morphology of organelles such as mitochondria and lysosomes were observed under an electron microscope. NADPH oxidase activity was observed in cells obtained from wild-type mice stimulated with PMA (Fig. 6). The reaction product was found in intracellular compartments of different sizes. No enzymatic reaction was observed in unstimulated cells, in PMA-stimulated cells isolated from wild-type mice incubated in a reaction medium containing 0.2 mM p-benzoquinone (Simic 1973), which is a scavenger of O2

–, or in PMA-stimulated cells isolated from gp91-phox-deficient mice (data not shown). Mitochondria retained their normal structure, and exhibited no swelling in PMA-stimulated cells positive for O2

– production. Lysosomes also remained structurally intact in the O2–-producing cells (Fig. 6).

of 2310.1007/s00418-004-0674-0


Fig. 6 Electron micrograph showing the localization of O2–-producing sites in a macrophage stimulated

with PMA isolated from wild-type mice. The reaction product was observed in intracellular compartments of different sizes (closed arrows). Mitochondria (open double arrows) and lysosomes (open arrows) retained their normal structure. Bar 1 m

Discussion The present study demonstrates that ROS produced intracellularly by NADPH oxidase activity lower the MTP in peritoneal resident macrophages stimulated with PMA isolated from wild-type mice, and that, on the contrary, MTP is preserved in PMA-stimulated cells obtained from mice deficient of gp91-phox, a membrane component of the NADPH oxidase.

of 2310.1007/s00418-004-0674-0


The O2–-producing intracellular compartments were herein detected in the macrophages isolated

from mouse peritoneal cavity using a cytochemical approach previously developed in our laboratory to detect O2

–-producing intracellular compartments in human living neutrophils stimulated with PMA by fluorescence microscopy (Kobayashi et al. 2000). The reliability of this method was confirmed by the comparison of neutrophils isolated from normal human peripheral blood with those obtained from patients with chronic granulomatous disease lacking gp91-phox, which are not able to produce O2

– (Roos et al. 1996). In the present study O2– was visualized in

PMA-stimulated wild-type macrophages, but not in PMA-stimulated gp91-phox-deficient macrophages obtained from mice which were generated by targeted disruption of the gp91-phox locus in 129-SV murine embryonic stem cells (Pollock et al. 1995; Dinauer et al. 2001), confirming that the O2

– detected in the macrophages is a product of the NADPH oxidase activity. We were also able to recognize the existence of two subpopulations in the macrophages isolated from wild-type mice: namely, a subpopulation of cells that produces O2

–, and another which does not. Mouse peritoneal macrophages have been reported to show heterogeneity with respect to phospholipid molecular species (Akoh and Chapkin 1990), tumor necrosis factor production (Bradbury and Moreno 1993), cell size, and lectin binding and antigen expression, as a result of the different stages of macrophage maturation (DaMatta et al. 1995). Accordingly, the activation of NADPH oxidase in mouse macrophages perhaps could also be associated with cell maturation.

Mitochondria play an essential role in apoptosis, and integrate diverse apoptotic stimuli into a core death pathway (Green and Reed 1998). One of the phenomena which appears in the early stage of apoptosis is the decrease in MTP (Zamzami et al. 1995), followed by the release of cytochrome c which complexes with Apaf-1 and procaspase-9 forming the apoptosome, a direct activator of downstream effector caspases-3 and -7 (Li et al. 1997; Budihardjo et al. 1999). In the present study, the decrease in MTP in cells producing O2

– was demonstrated by fluorescence microscopy, and also by flow cytometry. These findings seem to indicate that the subpopulation of macrophages producing O2

– suffers apoptosis during stimulation of the cells with PMA.

Swelling and loss of MTP have been reported to be associated with the release of cytochrome c during apoptosis in rat liver (Salvi et al. 2003), but there is also indication that the release of cytochrome c occurs as well in intact and functionally active mitochondria (Gogvadze et al. 2004). We observed no mitochondrial swelling in O2

–-producing cells under the electron microscope, indicating that the loss of MTP is not associated with mitochondrial swelling. In agreement with Gogvadze et al. (2004), it is, thus, speculated that the release of cytochrome c may occur in morphologically intact mitochondria.

Catalase is known to prevent apoptosis in T lymphocytes (Wesch et al. 1998) and in human neutrophils (Gamberale et al. 1998; Aoshiba et al. 1999), and has been reported to effectively inhibit both lung cell apoptosis and the inflammatory response, whereas superoxide dismutase (O2

– scavenger) and deferoxamine (inhibitor of ·OH generation by Fenton-like reactions) have lesser effects (Wang et al. 2003). Moreover, pretreatment with catalase, but not superoxide dismutase, completely inhibits capsaicin-induced apoptosis by inhibiting phosphorylation of the

of 2310.1007/s00418-004-0674-0


Ser-15 residue of p53 in leukemic cell lines (Ito et al. 2004), suggesting that H2O2 plays a major

role in apoptosis. In the present study, no MTP loss was observed in O2–-producing cells injected

with catalase, indicating that H2O2 is probably an effector molecule which disrupts MTP in

mouse macrophages. It is, therefore, suggested that ROS, including H2O2, leak from the O2–-

producing intracellular compartments to the cytoplasm, reach mitochondria, and eventually diminish MTP.

Reactive oxygen species such as H2O2 have also been reported to damage lysosomes, an event which is followed by the release of lysosomal enzymes (Brunk and Svensson 1999; Li et al. 2000; Brunk et al. 2001). Lysosomes play multiple roles in various cellular oxidative activities such as the oxidative burst during cytotoxic killing, and it has been demonstrated that a subset of lysosomes suffers alkalinization due to the increase of intralysosomal O2

– and H2O2 produced by

NADPH oxidase activity in macrophages stimulated with PMA (Chen 2002). The effect of O2–

on the localization of lysosomes was, therefore, examined by fluorescence microscopy. No alteration was, however, recognized in the lysosomal localization in O2

–-producing cells as

compared to those in O2–-deficient cells stimulated with PMA. The electron microscopic study

further revealed that lysosomes remain intact in O2–-producing cells. Accordingly, it is suggested

that no functional or morphological changes occur in the lysosomes in cells stimulated with PMA under the present experimental conditions (the cutoff time of the cell stimulation was 30 min).

In conclusion, we here demonstrate that: (1) the localization of O2–-producing sites in mouse

peritoneal resident macrophages stimulated with PMA can be visualized under the fluorescence microscope, (2) regarding O2

– production, two subpopulations of macrophages exist, and (3) the

ROS, including H2O2, dismutated from the O2– produced by NADPH oxidase activity, reduce

the MTP of the macrophages.

Acknowledgements This work was partially supported by Grants-in-Aid for Scientific Research (C) 13670968 and 15591281 from the Japanese Ministry of Education, Science, Sports, and Culture to Y.O. and T.K.

References

Akoh CC, Chapkin RS (1990) Composition of mouse peritoneal macrophage phospholipid molecular species. Lipids 25:613–617

Aoshiba K, Yasui S, Hayashi M, Tamaoki J, Nagai A (1999) Role of p38-mitogen-activated protein kinase in spontaneous apoptosis of human neutrophils. J Immunol 162:1692–1700

of 2310.1007/s00418-004-0674-0


Arroyo A, Modriansky M, Serinkan FB, Bello RI, Matsura T, Jiang J, Tyurin VA, Tyurina YY, Fadeel B, Kagan VE (2002) NADPH oxidase-dependent oxidation and externalization of phosphatidylserine during apoptosis in Me2SO-differentiated HL-60 cells. Role in phagocytic clearance. J Biol Chem 277:49965–49975

Babior BM (1999) NADPH oxidase: an update. Blood 93:1464–1476

Babior BM (2004) NADPH oxidase. Curr Opin Immunol 16:42–47

Badwey JA, Robinson JM, Lazdins JK, Briggs RT, Karnovsky MJ, Karnovsky ML (1983) Comparative biochemical and cytochemical studies on superoxide and peroxide in mouse macrophages. J Cell Physiol 115:208–216

Bradbury MG, Moreno C (1993) Effect of lipoarabinomannan and mycobacteria on tumour necrosis factor production by different populations of murine macrophages. Clin Exp Immunol 94:57–63

Briggs RT, Drath DB, Karnovsky ML, Karnovsky MJ (1975) Localization of NADH oxidase on the surface of human polymorphonuclear leukocytes by a new cytochemical method. J Cell Biol 67:566–586

Brunk UT, Svensson I (1999) Oxidative stress, growth factor starvation and Fas activation may all cause apoptosis through lysosomal leak. Redox Rep 4:2–11

Brunk UT, Neuzil J, Eaton JW (2001) Lysosomal involvement in apoptosis. Redox Rep 6:91–97

Budihardjo I, Oliver H, Lutter M, Luo X, Wang X (1999) Biochemical pathways of caspase activation during apoptosis. Annu Rev Cell Dev Biol 15:269–290

Chauhan D, Li G, Sattler M, Podar K, Mitsiades C, Mitsiades N, Munshi N, Hideshima T, Anderson KC (2003) Superoxide-dependent and -independent mitochondrial signaling during apoptosis in multiple myeloma cells. Oncogene 22:6296–6300

Chen CS (2002) Phorbol ester induces elevated oxidative activity and alkalization in a subset of lysosomes. BMC Cell Biol 3:21

of 2310.1007/s00418-004-0674-0


Chung YM, Bae YS, Lee SY (2003) Molecular ordering of ROS production, mitochondrial changes, and caspase activation during sodium salicylate-induced apoptosis. Free Radic Biol Med 15:434–442

DaMatta RA, Araujo-Jorge T, de Souza W (1995) Subpopulations of mouse resident peritoneal macrophages fractionated on Percoll gradients show differences in cell size, lectin binding and antigen expression suggestive of different stages of maturation. Tissue Cell 27:505–513

Dinauer MC, Gifford MA, Pech N, Li LL, Emshwiller P (2001) Variable correction of host defense following gene transfer and bone marrow transplantation in murine X-linked chronic granulomatous disease. Blood 97:3738–3745

Fadeel B, Ahlin A, Henter JI, Orrenius S, Hampton MB (1998) Involvement of caspases in neutrophil apoptosis: regulation by reactive oxygen species. Blood 92:4808–4818

Fratti RA, Backer JM, Gruenberg J, Corvera S, Deretic V (2001) Role of phosphatidylinositol 3-kinase and rab5 effectors in phagosomal biogenesis and mycobacterial phagosome maturation arrest. J Cell Biol 154:631–644

Gamberale R, Giordano M, Trevani AS, Andonegui G, Geffner JR (1998) Modulation of human neutrophil apoptosis by immune complexes. J Immunol 161:3666–3674

Gogvadze V, Robertson JD, Enoksson M, Zhivotovsky B, Orrenius S (2004) Mitochondrial cytochrome c release may occur by volume-dependent mechanisms not involving permeability transition. Biochem J 378:213–217

Green DR, Reed JC (1998) Mitochondria and apoptosis. Science 281:1309–1312

Hancock JT, Desikan R, Neill SJ (2001) Does the redox status of cytochrome c act as a fail-safe mechanism in the regulation of programmed cell death? Free Radic Biol Med 31:697–703

Ito K, Nakazato T, Yamato K, Miyakawa Y, Yamada T, Hozumi N, Segawa K, Ikeda Y, Kizaki M (2004) Induction of apoptosis in leukemic cells by homovanillic acid derivative, capsaicin, through oxidative stress: implication of phosphorylation of p53 at Ser-15 residue by reactive oxygen species. Cancer Res 64:1071–1078

Karnovsky MJ (1994) Cytochemistry and reactive oxygen species: a retrospective. Histochemistry 102:15–27

of 2310.1007/s00418-004-0674-0


Kobayashi T, Robinson JM (1991) A novel intracellular compartment with unusual secretory properties in human neutrophils. J Cell Biol 113:743–756

Kobayashi T, Seguchi H (1999) Novel insight into current models of NADPH oxidase regulation, assembly and localization in human polymorphonuclear leukocytes. Histol Histopathol 14:1295–1308

Kobayashi T, Seguchi H (2001) Cytochemistry of NADPH oxidase activity in human neutrophils. Acta Histochem Cytochem 34:85–89

Kobayashi T, Robinson JM, Seguchi H (1998) Identification of intracellular sites of superoxide production in stimulated neutrophils. J Cell Sci 111:81–91

Kobayashi T, Garcia del Saz E, Hendry J, Seguchi H (1999) Detection of oxidant producing-sites in glutaraldehyde-fixed human neutrophils and eosinophils stimulated with phorbol myristate acetate. Histochem J 31:181–194

Kobayashi T, Zinchuk VS, Okada T, Wakiguchi H, Kurashige T, Takatsuji H, Seguchi H (2000) A simple approach for the analysis of intracellular movement of oxidant-producing intracellular compartments in living neutrophils. Histochem Cell Biol 113:251–257

Kobayashi T, Tsunawaki S, Seguchi H (2001) Evaluation of the process for superoxide production by NADPH oxidase in human neutrophils: evidence for cytoplasmic superoxide. Redox Rep 6:27–36

Leavey PJ, Gonzalez-Aller C, Thurman G, Kleinberg M, Rinckel L, Ambruso DW, Freeman S, Kuypers FA, Ambruso DR (2002) A 29-kDa protein associated with p67phox expresses both peroxiredoxin and phospholipase A2 activity and enhances superoxide anion production by a cell-free system of NADPH oxidase activity. J Biol Chem 277:45181–45187

Li P, Nijhawan D, Budihardjo I, Srinivasula SM, Ahmad M, Alnemri ES, Wang X (1997) Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade. Cell 91:479–489

Li W, Yuan X, Nordgren G, Dalen H, Dubowchik GM, Firestone RA, Brunk UT (2000) Induction of cell death by the lysosomotropic detergent MSDH. FEBS Lett 470:35–39

Nishizuka Y (1986) Studies and perspectives of protein kinase C. Science 233:305–312

of 2310.1007/s00418-004-0674-0


Ogawa Y, Nishioka A, Kobayashi T, Kariya S, Hashimoto S, Saibara T, Nakayama K, Suguchi H, Yoshida S (2002) Mitochondrial cytochrome c release in radiation-induced apoptosis of human peripheral T cells. Int J Mol Med 10:263–268

Ogawa Y, Kobayashi T, Nishioka A, Kariya S, Hashimoto S, Seguchi H, Yoshida S (2003a) Radiation-induced oxidative DNA damage, 8-oxoguanine, in human peripheral T cells. Int J Mol Med 11:27–32

Ogawa Y, Kobayashi T, Nishioka A, Kariya S, Hashimoto S, Seguchi H, Yoshida Y (2003b) Radiation-induced reactive oxygen species formation prior to oxidative DNA damage in human peripheral T cells. Int J Mol Med 11:149–152

Ogawa Y, Kobayashi T, Nishioka A, Kariya S, Ohnishi T, Hamasato S, Seguchi H, Yoshida S (2004) Reactive oxygen species-producing site in hydrogen peroxide-induced apoptosis of human peripheral T cells: involvement of lysosomal membrane destabilization. Int J Mol Med 13:383–388

Pollock JP, Williams DA, Gifford MA, Li LL, Du X, Fisherman J, Orkin SH, Doerschule CM, Dinauer MC (1995) Mouse model of X-linked chronic granulomatous disease, an inherited defect in phagocyte superoxide production. Nat Genet 9:202–209

Robinson JM, Badwey JA (1995) The NADPH oxidase complex of phagocytic leukocytes: a biochemical and cytochemical view. Histochem Cell Biol 103:163–180

Robinson JP, Bruner LH, Bassoe CF, Hudson JL, Ward PA, Phan SH (1988) Measurement of intracellular fluorescence of human monocytes relative to oxidative metabolism. J Leukoc Biol 43:304–310

Roos D, de Boer M, Kuribayashi F, Meischl C, Weening RS, Segal AW, Ahlin A, Nemet K, Hossle JP, Bernatowska-Matuszkiewicz E, Middleton-Price H (1996) Mutations in the X-linked and autosomal recessive forms of chronic granulomatous disease. Blood 87:1663–1681

Salvi M, Fiore C, Armanini D, Toninello A (2003) Glycyrrhetinic acid-induced permeability transition in rat liver mitochondria. Biochem Pharmacol 66:2375–2379

Sauer H, Wartenberg M, Hescheler J (2001) Reactive oxygen species as intracellular messengers during cell growth and differentiation. Cell Physiol Biochem 11:173–186

Seguchi H, Kobayashi T (2002) Study of NADPH oxidase-activated sites in human neutrophils. J

of 2310.1007/s00418-004-0674-0


Electron Microscopy 51:87–91

Simic M (1973) Comparison between the electron transfer reactions from free radicals and their corresponding peroxy radicals to quinines. Biochem Biophys Res Commun 50:364–369

Spurr AR (1969) A low-viscosity epoxy resin embedding medium for electron microscopy. J Ultrastruct Res 26:31–43

Wadia JS, Chalmers-Redman RME, Ju WJH, Carlile GW, Phillips JL, Fraser AD, Tatton WG (1998) Mitochondrial membrane potential and nuclear changes in apoptosis caused by serum and nerve growth factor withdrawal: time course and modification by (-)-deprenyl. J Neurosci 18:932–947

Wang L, Medan D, Mercer R, Overmiller D, Leornard S, Castranova V, Shi X, Ding M, Huang C, Rojanasakul Y (2003) Vanadium-induced apoptosis and pulmonary inflammation in mice: role of reactive oxygen species. J Cell Physiol 195:99–107

Wesch D, Marx S, Kabelitz D (1998) Monocyte-dependent death of freshly isolated T lymphocytes: induction by phorbol ester and mitogens and differential effects of catalase. J Immunol 161:1248–1256

Xiao L, Pimentel DR, Wang J, Singh K, Colucci WS, Sawyer DB (2002) Role of reactive oxygen species and NAD(P)H oxidase in alpha(1)-adrenoceptor signaling in adult cardiac myocytes. Am J Physiol 282:C926–C934

Zamzami N, Marchetti P, Castedo M, Zanin C, Vayssiere JL, Petit PX, Kroemer G (1995) Reduction in mitochondrial potential constitutes an early irreversible step of programmed lymphocyte death in vivo. J Exp Med 181:1661–1672

of 2310.1007/s00418-004-0674-0


mitochondrial transmembrane potential is diminished in ... transmembrane potential.pdfmitochondrial...

Documents