Biol. Bull. 209: 168-183. (December 2005)
© 2005 Marine Biological Laboratory
Hydrogen Peroxide Induces Apoptotic-Like Cell Death in Coelomocytes of Themiste petricola (Sipuncula)
Guillermo A. Blanco1,*,¶,
Juanita Bustamante2,¶,
Mariana Garcia1,# and
Silvia E. Hajos1,¶
1 Department of Immunology, IDEHU-National Research Council (CONICET), Buenos Aires, Argentina
2 Department of Physicochemistry, School of Pharmacy and Biochemistry, University of Buenos Aires (UBA), Buenos Aires, Argentina
* To whom correspondence should be addressed. E-mail: gblanco{at}ffyb.uba.ar
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Abstract
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Apoptosis is an active form of cell death that plays a critical role in physiological and pathological conditions of multicellular organisms. These conditions include development, organogenesis, and elimination of infected, mutated, or damaged cells. Sipunculan cells may respond to changes in environmental exposure to oxidative stress by induction of apoptotic cell death. In coelomocytes of the sipunculan worm Themiste petricola, we evaluated morphological and biochemical changes that were induced by hydrogen peroxide (H2O2) and that could be compatible with an apoptotic-like phenotype. At an exposure of 100 mM H2O2, coelomocytes exhibited several morphological hallmarks of apoptosis such as chromatin condensation, nuclear segmentation, cell volume decrease, membrane blebbing, and formation of apoptotic bodies. Biochemical evidences of apoptotic-like cell death included exposure of phosphatidylserine (PS) in the outer leaflet of the plasma membrane and oligonucleosomal DNA fragmentation. In addition, exposure of coelomocytes to H2O2 induced a rapid massive loss of mitochondrial membrane potential and of the acidic pH of lysosomes. Overall, our results showed that, in sipunculan coelomocytes, H2O2 can induce changes compatible with an apoptotic-like phenotype. The finding of an oxidative-stress-induced apoptotic-like phenotype in a sipunculan worm may indicate that this kind of cell death process participates in regulation of cell number during physiological and pathological situations, including immune responses.
Abbreviations: AO, acridine orange • EB, ethidium bromide • JC-1, 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolcarbocyanine iodide • 
m, mitochondrial transmembrane potential • PI, propidium iodide • PS, phosphatidylserine
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Introduction
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Apoptosis is an active form of cell death with a vital role in physiological and pathological conditions throughout the development and adult life of multicellular organisms, removing infected, injured, or mutated cells (Kerr et al., 1972; Wyllie et al., 1980; Ellis and Horvitz, 1986). It is characterized by a series of morphological events such as cell shrinkage, chromatin condensation, nuclear and cytoplasmic fragmentation, and formation of dense bodies that are rapidly phagocytosed by neighboring cells (Vaux and Strasser, 1996). These typical morphological changes result from the activation of biochemical events such as increased intracellular Ca++ concentrations, uncoupling of mitochondrial electrical potential, fragmentation of oligonucleosomal DNA, and proteolytic degradation of specific substrates (Wyllie, 1980; Schwartzman and Cidlowski, 1993; Zheng et al., 1998; Enari et al., 1998). It is generally assumed that apoptosis appeared early in the evolution of multicellular animals, and several studies have shown that the components of the cell death pathway are highly conserved throughout the evolution of Caenorhabditis elegans, Drosophila, and mammals (Meier et al., 2000; Vernooy et al., 2000).
Apoptosis can be induced under several stress conditions such as withdrawal of growth factor (Xia et al., 1995), exposure to cytotoxic drugs (Muschel et al., 1998; Lopes et al., 2003), or loss of normal adhesion to extracellular matrix (Frisch and Francis, 1994). Environmental pollutants such as heavy metals and organotines are known to induce apoptosis in invertebrates and vertebrates and may thus incline the equilibrium of cellular homeostasis towards an increased cellular mortality (Micic et al., 2001; Sokolova et al., 2004). Enhanced cell death in the hemocyte population of marine invertebrates has the potential to generate an immunosuppressed organism with a reduced capacity to resist pathological insults and opportunistic infections. There are few publications documenting apoptosis in marine invertebrates, especially in relation to the morphological changes of the cells. The effects of temperature and salinity on apoptosis have been studied in the hemocytes of Crassostrea virginica after in vitro incubation (Goedken et al., 2005a). Parasite infection may also influence apoptotic balance since in vitro infection with Perkinsus marinus suppressed apoptosis in C. virginica hemocytes (Goedken et al., 2005b). It has been suggested that the parasite may inhibit hemocyte apoptosis to provide more host cells for itself (Sunila and LaBanca, 2003; Goedken et al., 2005b). Thus, an understanding of apoptosis at a cellular and molecular level is important because excessive apoptosis of hemocytes may be detrimental to the ability of marine invertebrates to fight infectious diseases (Sunila and LaBanca, 2003).
Morphological changes, such as nuclear and chromatin condensation and fragmentation, that occur in apoptosis can be assessed by fluorescent microscopy using supravital dyes such as acridine orange (AO) and ethidium bromide (EB). Used together, these dyes can discriminate between viable and nonviable cells and allow the identification of morphological changes that occur during apoptosis (Lopes et al., 2003). Biochemical hallmarks of apoptosis such as translocation of phosphatidylserine (PS) to the extracellular side of the plasma membrane and fragmentation of oligonucleosomal DNA are best detected, respectively, by Annexin-V FITC binding and agarose gel electrophoresis. All the above-mentioned morphological and biochemical changes uncompromisingly indicate apoptosis, especially when evaluated together. Blebbing and volume decrease may be considered part of the apoptotic phenotype when present in conjunction with the aforementioned morphological and biochemical apoptotic changes. In addition, the reduction of mitochondrial transmembrane potential (m) is considered an apoptotic hallmark in higher vertebrates (Kroemer et al., 1997), although a recent study in oyster hemocytes demonstrated that apoptosis may occur without loss of m (Sokolova et al., 2004). Changes in m can be evaluated by fluorescent probes such as 5,5',6,6'-tetrachlorol, 1',3,3'-tetraethylbenzimidazolcarbocyanine iodide (JC-1) (Cossarizza et al., 1995; Kulkarni et al., 1998).
Sipunculans are coelomate nonsegmented marine worms closely related to annelids; however, they lack a true circulatory system (Stephen and Edmonds, 1972; Maxmen et al., 2003). The coelomic cavity of sipunculans contains a number of cell types. Hemerythrocytes constitute about 90% of the coelomocytes (Rice, 1993). They are 10–20 µm diameter, bear the respiratory pigment hemerythrin, are more or less biconvex in shape, and have large lysosomic vacuoles that are variable in number depending on the species (Ochi, 1975). Species of the genus Themiste characteristically have a single lysosomic vacuole (Ochi, 1975). Amebocytes are the second most abundant cell type and are more variable in form and structure both within and among species than are hemerythrocytes (Rice, 1993). Types of amebocytes, not necessarily all present in the same species, include hyaline forms with fine or no granules and granulocytes with coarse granules (Hyman, 1959; Rice, 1993). Hyaline amebocytes are phagocytic and, like hemerythrocytes, have large lysosomic vacuoles (Ochi, 1975). Various types of multicellular bodies, such as urn cells—large multinuclear cells or "granulocytes with an associated cell"—are reported in different species (Hyman, 1959; Dybas, 1975, 1981; Rice, 1993). Sipunculans are dioecious, and either ovocytes in females or clumps of spermatozoa in males can be seen in the coelomic fluid (Hyman, 1959; Rice, 1993).
Coelomocytes from sipunculid worms have been shown to participate in immune responses such as phagocytosis and cytotoxicity against foreign agents (Boiledieu and Valembois, 1977; Blanco et al., 1995, 1997; Cabrera et al., 2002). Given the importance of a homeostatic balance of apoptosis in immunity, regulation of coelomocyte apoptosis could be significant to the sipunculan immune system, and several environmental factors could have an impact on the coelomocytes, thus triggering the apoptotic program. Hydrogen peroxide (H2O2) can produce apoptosis in the cells of most animals from protozoa to mammals (Palomba et al., 1999; Li et al., 2000; Das et al., 2001). H2O2 is considered to induce apoptosis through a massive depolarization of mitochondria (Marzo et al., 1998).
The aim of this study was to evaluate changes that could be considered compatible with an ongoing apoptotic-like process in coelomocytes of the sipunculan Themiste petricola Amor, 1964, after exposure to H2O2. We evaluated nuclear and cytoplasmic morphological changes, mitochondrial membrane-potential changes, and the presence of DNA fragmentation, as well as translocation of PS to the extracellular side of the plasma membrane as detected by Annexin-V binding. Our results were consistent with the induction of an apoptotic phenotype as a result of coelomocyte exposure to H2O2.
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Materials and Methods
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Coelomocyte isolation, primary culture, and hydrogen peroxide exposure
Adult specimens of Themiste petricola with an average wet weight of 100 mg were collected from crevices in intertidal rocks at Santa Elena beach on the coast of Argentina, 34 °S latitude. Animals were maintained in 500-ml plastic boxes with frequently renewed filtered seawater (3.2% salinity) at 18 °C. Coelomocyte suspensions were prepared as indicated previously (Blanco et al., 1995). In brief, coelomic fluid was harvested by incision of the body wall with a sterile surgical blade, allowing the fluid to drip into a 15-ml sterile centrifuge tube cooled to 4 °C. The suspension was centrifuged at 300 x g, the supernatant was discarded, and the coelomocyte pellet was washed three times by centrifugation at 300 x g in balanced salt solution (BSS) made to resemble the ionic composition of T. petricola coelomic fluid. The composition of BSS was 0.4 M NaCl, 10.3 mM KCl, 10.1 mM CaCl2, 30.2 mM MgS04, 26.2 mM NaHCO3, pH 7.0. Germinal cells were excluded with sterile nylon linen (mesh: 50 µm). For in vitro assays, coelomocytes were suspended in RPM1 1640 medium supplemented with NaCl, KCl, CaCl2, MgSO4, and NaHCO3, as required to achieve the same concentrations cited for BSS. Coelomocyte suspensions of 1 x 106 cells in 1 ml RPMI in Eppendorf tubes were exposed to 100 mM H2O2 and assayed for different apoptotic indicators at several time points. Control non-exposed coelomocyte suspensions were incubated under similar conditions and assayed to detect spontaneous apoptosis.
Fluorescence microscopy
All fluorescence observations were made with a Zeiss Standard 14 photomicroscope with epifluorescence system equipped with an automatic C 35 camera loaded either with Kodak Gold 100 ASA film, for slides stained with acridine orange (AO) and ethidium bromide (EB), or with Kodak Ultra 400 ASA, for slides strained with JC-1 and Annexin V-FITC. Film negatives were digitized with a high-resolution film scanner (Kodak Professional Film Scanner at 3600 dpi). Digital images were processed using the public domain Image J (NIH, Bethesda, MD) image analysis software.
Phagocytosis assay
Coelomocytes (1 x 106) were incubated for 30 min at 20 °C with NH4Cl—10 mM, 50 mM, and 100 mM—in 1 ml RPMI, in Eppendorf tubes. In a first series of experiments, cells were centrifuged and further resuspended in 25 µl of RPMI and 2 µl of AO/EB dye mixture to evaluate changes in AO fluorescence due to elevated intra-lysosomal pH. AO is an optical probe used to study vacuolar acidification because it shows different absorption and fluorescence properties in its different forms (Millot et al., 1997). Since AO is a lipophilic weak base that is membrane-permeable in its neutral form but relatively membrane-impermeable once protonated, it becomes trapped in acidic vesicles, where high concentrations of the protonated form are achieved (Barasch et al., 1991). Thus, low-pH organelles are stained red by AO due to high concentrations driven by the pH gradient through membrane vesicles, while neutral or high-pH organelles are stained green. Agents such as NH4Cl are relatively lipophilic in their un-protonated form and pass through the membranes of vacuoles. In an acidic environment, these molecules become protonated and cannot escape from these organelles, which favors their accumulation and the increase in vacuolar pH.
In a second series of experiments, both NH4Cl-treated cells and control non-treated cells were resuspended in 1 ml of RPMI containing 1% (w/v) zymosan (SIGMA, St. Louis) and were further incubated for 40 min to allow phagocytosis to proceed. Smears were prepared with aliquots from each tube and were fixed in May Grunwald and stained with 10% Giemsa to morphologically assess phagocytosis of zymosan particles by NH4Cl-treated coelomocytes and control non-treated coelomocytes. In other series of phagocytosis experiments performed in the same conditions, coelomocytes were suspended in 25 µl of RPMI 1640 and 2 µl of a mixture of AO and EB dyes (AO/EB) and immediately examined under fluorescence microscopy.
Apoptosis indicator staining
Acridine orange and ethidium bromide (AO/EB) staining.
After different times of exposure with H2O2, coelomocyte pellets with 1 x 106 cells were suspended in 25 µl of RPMI 1640 and 2 µl of AO/EB dye mix (100 µg ml–1 acridine orange plus 100 µg ml–1 ethidium bromide in PBS) and immediately examined under fluorescence microscopy (Lopes et al., 2003). The time course of apoptosis was evaluated by scoring a minimum of 200 cells and determining the number of cells in each of three groups: apoptotic viable, apoptotic dead, and non-apoptotic viable.
Measurement of mitochondrial transmembrane potential.
m was measured using the mitochondria-specific fluorescence probe JC-1 (Molecular Probes, Eugene, OR). Stock solution of JC-1 was prepared at 5 mM in dimethylsulfoxide. This dye is incorporated into mitochondria and experiences a shift in fluorescence emission spectra when the membrane is depolarized (Cossarizza et al., 1995; Kulkarni et al., 1998). Suspensions of either H2O2-exposed or control non-exposed coelomocytes (1 x 106 in 250 µl of RPMI) were incubated in 5 µM JC-1 and observed under fluorescence microscopy with a 100x oil immersion objective. A second series of experiments was performed under similar conditions and evaluated by flow cytometry.
Annexin V-FITC labeling and flow cytometry.
The expression of phosphatidylserine (PS) in the outer membrane of H2O2-exposed coelomocytes and control non-exposed coelomocytes was monitored by labeling with Annexin V-FITC (Biovision, Mountain View, CA) and evaluated by flow cytometry and fluorescence microscopy. Coelomocytes were washed once in 1 ml of BSS, resuspended in 20 µl of binding buffer (10 mM HEPES, 0.4 M NaCl, 2.5 mM CaCl2) with 2.5 µl of Annexin V-FITC, and incubated for 10 min. For fluorescence microscopy, 10-µl aliquots were placed on microscope slides and observed with a 100x oil immersion objective. For flow cytometry, coelomocytes were suspended in 500 µl of binding buffer. Propidium iodide (PI) was added prior to observation by flow cytometry or fluorescence microscopy to further discriminate between PS exposure in viable coelomocytes (early apoptotic, Annexin V-FITC-positive only) and nonviable coelomocytes (late apoptotic, Annexin V-FITC-positive and PI-positive due to DNA labeling).
A FACScalibur (Beckton Dickinson, San Jose, CA) flow cytometer was used to detect Annexin V-FITC and PI fluorescence. The excitation wavelength was 488 nm for both FITC and PI; for detection, 530-nm (FITC) and 585-nm (PI) band pass filters were used. Fluorescence parameters from single cells were collected using a logarithmic amplifier after gating on the combination of forward light scatter (FSC) and perpendicular light scatter (SSC). Red fluorescence from PI was collected through the FL2 channel, and green fluorescence from Annexin V-FITC through the FL1 channel. For studies of JC-1 supravital staining, the excitation wavelength was 488 nm; 530-nm band pass filters were used to detect green fluorescence (JC-1 monomers), and 585-nm band pass filters were used for orange fluorescence (JC-1 aggregates). A total of 10,000 cells were analyzed per tube, and data acquired in list mode were processed using WinMDI 2.8 software (J. Trotter, The Scripps Research Institute, La Jolla, CA). The fluorescence distribution was displayed as a two-color dot plot analysis, and the percentage of fluorescent cells in each quadrant was determined.
Assessment of nuclear DNA oligonucleosomal fragmentation.
Coelomocyte pellets with 5 x 106 cells were lysed with 500 µl of lysis buffer (500 mM Tris-HCl pH 7.5, 1 mM EDTA, 0.2% Triton X-100) centrifuged at 13,000 x g for 10 min, and the supernatant containing the fragmented DNA was precipitated overnight at –20 °C in 700 µl of isopropanol and 100 µl of 5 M NaCl. The pellet was collected after centrifugation at 13,000 x \E g for 10 min, air-dried, and suspended in 10 mM Tris-HCl, 1 mM EDTA pH 7.5. A loading buffer containing 15 mM EDTA, 2% SDS, 50% glycerol, and 0.05% bromophenol blue was added at 1:5 (v/v). Samples were electrophoresed in 1% agarose gel and stained with 0.5 µg ml–1 ethidium bromide; DNA was visualized using ultraviolet light (Lopes et al., 2003). The DNA molecular weight marker was from PB-L Productos Bio-Logicos, UNQ, Argentina. Cells from a murine lymphoid leukemia (Lopes et al., 2003) were used as control and were assessed for DNA fragmentation in parallel to coelomocytes under the same protocol. Gel images were obtained with a digital camera (Olympus, Camedia, D-510).
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Results
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The normal appearance of sipunculan coelomocytes supravitally stained with acridine orange/ethidium bromide
Supravital staining with AO/EB revealed the presence of large vacuoles and granules in a large proportion of sipunculan coelomocytes. Viable large coelomocytes having one or a few large red vacuoles and a green nucleus were often seen (Fig. 1a). These large vacuoles could be identified as a clear area of cytoplasm on smears of coelomocytes stained with Giemsa (Fig. 1b). Under phase contrast microscopy, vacuoles often appeared to be of a dense material and were readily distinguishable in the cytoplasm (Fig. 1c).
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Figure 1. Untreated coelomocytes supravitally stained with acridine orange/ethidium bromide. Arrows point to nuclei exhibiting green fluorescence, shown here in light gray; arrowheads point to vacuoles exhibiting red fluorescence, shown here in dark grey. (Magnification 1000x; scale bars = 15 µm.) (A) Viable coelomocytes with green fluorescent nuclei and large red fluorescent vacuoles in proximity to the nuclei. (B) Coelomocytes stained with Giemsa also had large vacuoles. (C) Combined fluorescent and phase contrast microscopic view of viable coelomocytes.
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Acridine orange as a probe for acidic organelles of coelomocytes
AO becomes concentrated within acidic organelles such as lysosomes, causing a change in fluorescence from green to red (Millot et al., 1997). To test whether a low pH of vacuoles and granules was causing the accumulation of the dye and the metachromatic effect, we exposed cells to three doses of ammonium chloride for 1 h. Like AO, ammonium chloride becomes trapped and concentrated in lysosomes as a consequence of low pH, but its accumulation raises the pH of the lysosomes. At a 1-mM dose of ammonium chloride, granules and vacuoles were still stained red, indicating that the pH change was not enough to block the metachromatic effect of AO (Fig. 2a); at 50 mM, red granules and vacuoles were almost absent (Fig. 2b); at 100 mM, no red staining of granules or vacuoles was observed (Fig. 2c). The green staining of the nucleus indicated that, except for having an elevated pH in granules and vacuoles, the viability of coelomocytes was not affected.
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Figure 2. Coelomocytes exposed to NH4Cl supravitally stained with acridine orange/ethidium bromide. Arrows point to nuclei exhibiting green fluorescence, shown here in light gray; arrowheads point to vacuoles exhibiting red fluorescence, shown here in dark grey. (A) Coelomocytes exposed to 1 mM NH4Cl for 1 h show green fluorescent nuclei and large red fluorescent vacuoles in proximity to the nuclei. The presence of fluorescent vacuoles indicates that the pH change at this concentration of NH4Cl was not sufficient to block the metachromatic effect of AO/EB. (Magnification 250x; scale bar = 50 µm.) (B) Coelomocytes exposed to 50 mM NH4Cl for 1 h show green fluorescent nuclei and large red fluorescent vacuoles in proximity to the nuclei. Vacuoles were fewer and smaller under this treatment than in A. (Magnification 250x; scale bar = 50 µm). (C) Coelomocytes exposed to 100 mM NH4Cl for 1 h show green fluorescent nuclei; vacuoles were not detected under this treatment since red fluorescence was completely eliminated. The darker gray area surrounding the nuclei corresponds to green fluorescence all over the cytoplasm. (Magnification 400x; scale bar = 30 µm.) (D) Coelomocytes exposed to 100 mM NH4Cl were able to phagocytose zymosan particles (arrows), but most particles remained undegraded within the cytoplasm. Giemsa stain. (Original magnification 1000x, digitally enlarged; scale bar = 15 µm.) (E) Control non-exposed coelomocytes engulfed zymosan particles (arrows) and rapidly degraded them within the cytoplasm. Giemsa stain. (Original magnification 1000x, digitally enlarged; scale bar = 15 µm.) (F) Coelomocytes exposed at 100 mM NH4Cl showed undegraded zymosan particles (arrows) all over the cytoplasm when supravitally stained with AO/EB. All gray tones correspond to green fluorescence. (Original magnification 1000x, digitally enlarged; scale bar = 15 µm.) (G) Supravital staining with AO/EB of control non-exposed coelomocytes showed that zymosan particles (arrows) were rapidly degraded within the cytoplasm. Light gray round areas correspond to green fluorescent nuclei; darker gray areas correspond to red fluorescent vacuoles. (Original magnification 1000x, digitally enlarged; scale bar = 15 µm.)
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An elevated pH of granules and vacuoles may preclude optimal activity of acidic degradative enzymes. Thus, we evaluated the effect of raising the pH of granules and vacuoles in coelomocyte endocytosis and degradation of zymosan particles. Coelomocytes exposed to 100 mM ammonium chloride were able to phagocytose zymosan particles but could not successfully degrade endocytosed particles. Phagocytic coelomocytes became densely packed with undegraded zymosan particles dispersed throughout the cytoplasm (Fig. 2d and f). In contrast, control non-exposed coelomocytes exhibited only a few undegraded particles in the cytoplasm (Fig. 2e and g). Most of these particles were in proximity to the plasma membrane and may have been in the initial stages of engulfment. Thus, it appears that engulfment of particles is followed by rapid degradation, probably within the vacuole.
Changes induced by in vitro exposure of coelomocytes to hydrogen peroxide
Within the first 30 min of exposure of coelomocytes to 100 mM H2O2, most cells were viable, as revealed by green staining of the nuclei. No changes in nuclear morphology were observed, but some cytoplasmic condensations became evident together with a sharp decrease in red fluorescence of the vacuoles (Fig. 3a). In addition, vacuoles were often seen in proximity to the cell membrane instead of in the more central location seen in normal non-exposed cells (Fig. 3a). After 1 h of H2O2 exposure, most cells remained viable but had completely lost red fluorescence of the vacuoles, indicating an impairment to preserve the pH gradient in these organelles; and many cells showed morphological changes in the nucleus. These changes included chromatin focal condensations and loss of the homogenous fluorescence (Fig. 3b). Nuclear contour changed from round to irregular, and nuclear membrane ripples could occasionally be seen. Several patterns of chromatin distribution and nuclear shape alterations were observed, including marginal condensation, multiple small condensations, central condensations, and rippled contour of the nucleus (Fig. 3c). With longer incubation, these nuclear morphological changes seen after 1 h of H2O2 exposure became evident in many more cells. An overall decrease in the size of viable cells was seen beyond 1 h of H2O2 exposure; however, not all cells decreased their volume at the same pace, and a range of cytoplasmic sizes could be detected in the same microscope field (Fig. 3d and e). Cells with maximal shrinkage ultimately underwent extensive blebbing and cellular fragmentation, forming apoptotic bodies.
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Figure 3. Coelomocytes exposed to 100 mM H2O2 supravitally stained with acridine orange/ethidium bromide. (A) After 30 min of exposure to H2O2, most cells remained viable. Arrows point to light gray areas corresponding to green fluorescent nuclei. No changes were observed in nuclear shape, and the density of the chromatin was homogenous. Arrowheads point to darker gray areas corresponding to vacuoles having decreased red fluorescence that were often seen in proximity to the cell membrane rather than in the more central location observed in normal non-exposed cells. Cell size was preserved, but cytoplasmic condensations became evident. (Magnification 1000x; scale bar = 15 µm.) (B) Coelomocytes after 1 h of H2O2 exposure. Cells exhibited nuclear morphological changes including focal chromatin condensations, irregular shape, and occasionally nuclear membrane ripples. Arrows point to heterogeneous gray areas corresponding to green fluorescent nuclei. There was a complete loss of fluorescence from vacuoles. All gray tones in the panel correspond to green fluorescence. (Magnification 1000x; scale bar = 15 µm.) (C) A possible sequence of nuclear morphological changes as inferred from observations after a 1-h incubation of coelomocytes with 100 mM H2O2. (Original magnification 1000x; sample nuclei were digitally cropped from images and enlarged.) (D) Decreased cell size together with an increase in granularity in coelomocytes exposed for 1 h to H2O2 (arrows). (Phase contrast microscopy at magnification of 1000x; scale bar = 15 µm.) (E) A possible sequence of cytoplasmic changes and cell volume decrease, as inferred from observations after 1 h of incubation with H2O2. (Original magnification 1000x, samples digitally cropped from images and enlarged.)
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Control coelomocytes not exposed to H2O2 exhibited JC-1 orange fluorescence due to the formation of JC-1 aggregates, indicating that the mitochondria were polarized (Fig. 4a and b). The JC-1 accumulation and the orange fluorescence observed in normal mitochondria were due to the asymmetric distribution of protons and other ions on both sides of the inner mitochondrial membrane (Attardi and Schatz, 1988). Orange fluorescent mitochondria seemed to be motile and often appeared as large spots or in the shape of filaments (Fig. 4a and b). Exposure to H2O2 induced a very rapid decline in m, as revealed by complete loss of JC-1 orange fluorescence and a shift to JC-1 green fluorescence due to formation of JC-1 monomeres within the first 30 min (Fig. 4c and d). When evaluated by flow cytometry, control non-exposed coelomocytes exhibited a predominantly orange-green fluorescence (Fig. 5a). Coelomocytes exposed to H2O2 showed a time-dependent reduction in m, as revealed by a progressive decrease of JC-1 orange-green fluorescence and a shift to JC-1 green-only fluorescence (Fig. 5b to d). Control non-exposed coelomocytes exhibited a predominance of orange-green fluorescence even when evaluated after incubation times of 3 h and 5 h, indicating that the cells maintained a stable m (Fig. 5e and f).
On the basis of the observed apoptotic-like phenotype in coelomocytes stained with AO/EB, we evaluated the time course of changes induced by H2O2 (Fig. 6a). At 2 h of H2O2 exposure, most cells had undergone morphological apoptotic-like changes, and a small proportion of dead cells were observed. From 2 h to 5 h, viable apoptotic-like coelomocytes were most abundant, while the number of dead cells continued to increase. A small number of viable cells with a non-apoptotic phenotype were observed at 2 h but these were almost undetectable after 4 h of incubation. Control non-exposed coelomocytes did not exhibit a significant number of cells with an apoptotic-like phenotype at any time point evaluated between 1 and 5 h (Fig. 6b).
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Figure 6. Time-course evaluations of control and experimental coelomocytes. (A) Morphological changes in coelomocytes exposed to 100 mM H2O2. Apoptotic viable group included cells with green nucleus undergoing shape changes and chromatin changes (see Fig. 3B) and decreases in cell size together with increases in granularity (see Fig. 3D). Non-apoptotic viable group included coelomocytes with green nucleus, red fluorescent vacuole, no increased granularity, and normal size (see Fig. 1C). Apoptotic dead group included coelomocytes with red nucleus like those at the end stage of the series (see Fig. 3C), together with decreased cell size and increased cytoplasmic granularity or blebbing with formation of apoptotic bodies. Mean values of the percentage of cells in each group at each time point are from three independent experiments. Bars indicate standard error of the mean. (B) Morphological evaluation of apoptosis in coelomocytes exposed to 100 mM H2O2 and in control non-exposed coelomocytes that were cultured in RPMI medium alone. Apoptotic cells included both apoptotic viable and apoptotic dead coelomocytes. Mean values of the percentage of cells in each group at each time point are from three independent experiments. Bars indicate standard error of the mean. (C) Oligonucleosomal DNA fragmentation. Electrophoresis of DNA in 1% agarose gel in TE buffer. Lane 1: DNA molecular weight marker. Lanes 2 to 7: non-exposed coelomocytes used as a control to determine spontaneous apoptosis, evaluated at 0 h, 1 h, 2 h, 4 h, 6 h, and 8 h, respectively. Lanes 8 to 12: coelomocytes exposed to 100 mM H2O2 and evaluated at 1 h, 2 h, 4 h, 6 h, and 8 h, respectively. Lane 13: murine cells from a lymphoid leukemia exposed to 100 mM H2O2 and evaluated at 8 h.
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Evaluation of oligonucleosomal DNA fragmentation
Coelomocytes exposed to 100 mM H2O2 and followed up for 8 h to evaluate oligonucleosomal DNA fragmentation exhibited a typical DNA laddering pattern. Fragments separated by an interval of about 200 base pairs became evident after 4 h of exposure and continued to increase until sampling was discontinued at 8 h (Fig. 6c, lanes 8 to 12). This laddering pattern was similar to that of murine cells that were exposed in parallel to similar concentrations of H2O2, although the number of base pairs in the intervals between fragments appeared to be slightly larger in coelomocytes than in murine cells (Fig. 6c, lane 13). Control non-exposed coelomocytes did not exhibit a DNA laddering pattern at any time point evaluated between 0 and 8 h (Fig. 6c, lanes 2 to 7).
Exposure of phosphatidylserine on the outer side of coelomocyte cytoplasmic membrane
Exposure to 100 mM H2O2 caused the appearance of PS on the outer side of the coelomocyte cytoplasmic membrane, as revealed by labeling with Annexin V-FITC. The binding was detected after 2 h of incubation and was still apparent at 5 h (Fig. 7c and d, respectively). An Annexin V-FITC positive control was introduced by exposing coelomocytes to formaldehyde, which caused PS to be exposed (Fig. 7b). The simultaneous labeling with propidium iodide showed an increase in double-positive cells, corresponding to late apoptotic cells, at 5 h of H2O2 incubation. The exposure of PS occurred in parallel to cell shrinkage, and most double-positive late apoptotic cells had decreased in size and increased in granularity (Fig. 4e and f). Control non-exposed coelomocytes were not labeled with Annexin V-FITC or PI at any time point evaluated between 1 h and 8 h (Fig. 7a and Fig. 7e to h).
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Figure 7. Dot plots of control and experimental coelomocytes. (A) Control non-exposed coelomocytes after 5-h incubation, followed by staining with Annexin V-FITC, showing non-apoptotic viable cells. (B) Positive control for Annexin V-FITC obtained by incubating coelomocytes for 1 h in BSS-formaldehyde 4%. (C) Coelomocytes exposed to 100 mM H2O2 for 2 h. (D) Coelomocytes exposed to 100 mM H2O2 for 5 h. (E to H) Control non-exposed coelomocytes incubated in RPMI medium for 1 h, 2 h, 5 h, and 8 h, respectively, and further stained with Annexin V-FITC and propidium iodide.
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Discussion
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Animal cells may respond to changes in environmental exposure to oxidative stress and to endogenous free radical production by increases or decreases in cell proliferation, changes in immune response, or induction of apoptotic cell death (Jackson et al., 2002). Extreme levels of oxidative stress may induce necrotic cell death, a pathological process that has been considered a passive process (Samali et al., 1999). The effects of oxidative stress exposure have been thought to depend on the cell type, the level of oxidative stress, and the protective antioxidant mechanisms in a given cell type (Banki et al., 1996). In our study, in vitro exposure of sipunculan coelomocytes to 100 mM H2O2 induced a series of morphological and biochemical changes that were consistent with the induction of an active program of cell death.
The procedure we followed to harvest coelomocytes excluded gametes and yielded a cell suspension consisting mainly of hemerythrocytes and hyaline amebocytes—both cell types characterized by a single lysosomic vacuole as in other species of Themiste (Ochi, 1975). Small granular amebocytes were rare and constituted a minor cell type in the coelomocyte suspensions obtained, while multicellular bodies were not observed at all. The prominent acidic vacuole of coelomocytes was supravitally stained red with a mixture of acridine orange and ethidium bromide (AO/EB). We were interested in identifying the physiological function of these large vacuoles, and therefore we determined intra-vacuole pH and its effect on zymosan breakdown within the vacuole. In quantitative terms, the degree of accumulation depends on the transmembrane pH gradient and on the total internal volume of the acidic vesicles.
Lysosomes have been defined as intracellular digestive organelles that are generally assumed to have an internal pH of 4.0–5.5 generated by proton pumps (Ohkuma and Poole, 1978; Yamashiro et al., 1983; Kornfeld and Mellman, 1989). This characterization of lysosomes has been supported by the fact that many lysosomal enzymes require an acidic pH for optimal activity (Bond and Butler, 1987). Using confocal microscopy and multidrug-resistant K-562 cells, Millot et al., (1997) estimated that organelles stained red with AO were in a pH range of 5.5–4. Thus lysosomes are the only organelles that acquire such a low pH, because their enzymes have an acidic pH optimum. In our study we observed that raising the pH of large vacuoles by exposure to NH4Cl prevented ingested zymosan particles from being degraded, suggesting that degradative enzymes may be present in the vacuole of coelomocytes and require an acidic pH for optimal activity. Thus the acidic vacuole of coelomocytes has several features that correspond to those that characterize lysosomes and may be the main degrading organelle of these cells.
Recent work strongly implicates lysosomes in the resealing response to damage associated with wounds to the cell cortex in nucleated animal cells. Protein markers of the lysosome membrane have been detected on the surface of cells at sites of disruption, and rapid resealing has been shown to involve a Ca2+-dependent exocytosis response (Reddy et al., 2001; McNeil, 2002; Andrews, 2002). At early stages following H2O2 exposure, we observed the red fluorescent vacuoles of coelomocytes in proximity to the cell membrane rather than in the more central location seen in normal non-exposed cells. In addition, large acidic vacuoles began to lose red fluorescence within the first 30 min of exposure, suggesting a failure in proton pump activity and a decreased pH gradient associated with damage to the plasma membrane from massive oxidative stress. The relocation of the lysosomic vacuoles could be an early exocytotic attempt to repair the damaged membrane by fusing the large vacuoles to it and creating, in effect, a "patch" (McNeil, 2002).
Along with lysosomes, we observed loss of mitochondrial transmembrane potential (m), as indicated by suppression of orange-green fluorescence and a shift to green-only fluorescence following exposure to the JC-1 probe. Oxidative stress induces decreased m by opening the permeability transition pore, a multiprotein complex built up at the contact site between the inner and outer mitochondrial membranes (Marzo et al., 1998). Current paradigms of apoptosis suggest that the loss of m occurs early in the commitment phase of apoptosis that results in the release of mitochondrial apoptogenic proteins including cytochrome c and the apoptotic inducing factor (Zamzani et al., 1995; Kluck et al., 1997). Interestingly, lysosomal membrane permeabilization triggers collapse of the m in an organelle-specific induction of apoptosis (Boya et al., 2003). This fact may relate our observation of early alterations in lysosomic vacuoles with the parallel collapse of m.
Apoptosis-associated nuclear condensation and segmentation are generally attributed to caspase activity that leads to the cleavage of substrates such as lamins, caspase-activated DNAse, or acinus (Rao et al., 1996; Liu et al., 1997; Sakahira et al., 1998; Sahara et al., 1999). We have identified a series of nuclear morphological changes in viable coelomocytes. These changes, which we began to notice after 1 h of exposure and continued to see in the following hours, included rippled nuclear contour, chromatin condensation, and nuclear segmentation, which are consistent with patterns of nuclear apoptosis. It is possible that these changes depend on caspase activation, as in other animal cells (Enari et al., 1998; Zheng et al., 1998). Nevertheless, the presence of oligonucleosomal fragmentation of DNA in H2O2-exposed coelomocytes is suggestive of a complete course of apoptosis, since it is a hallmark of the apoptotic phenotype and is considered a late apoptotic and caspase-dependent event.
In normal cells, phosphatidylserine (PS) is found predominantly in the inner layer of the plasma membrane bilayer. Apoptotic cells lose this phospholipid asymmetry, leading to the exposure of PS on their surface. Thus, the binding of Annexin V to H2O2-exposed coelomocytes indicates PS exposure on the outer cell membrane leaflet, which was identified in the early stages of exposure and was coincident with initial changes in nuclear chromatin. In other species, exposure of PS is thought to facilitate recognition of the cells as apoptotic, triggering phagocytosis by macrophages (Fadok et al., 1992). A ubiquitous and distinctive feature of apoptosis is the loss of cell volume, which is common to all examples of apoptosis, independent of cell type (Bortner and Cidlowski, 1998). It has recently become evident that the normotonic cell shrinkage starting before cell fragmentation, termed the apoptotic volume decrease (AVD) (Maeno et al., 2000), is induced by activation of ionic processes involving K+ and Cl– channels (Okada and Maeno, 2001). In contrast, necrosis is characterized by an increased cell volume (Bortner and Cilowski, 1998). We have identified AVD in coelomocytes exposed to H2O2 as an early event that progressed slowly along with changes in the nucleus and exposure of PS. Induction of AVD in other species precedes cytochrome c release, caspase-3 activation, and DNA laddering; and it is considered a caspase-independent event (Maeno et al., 2000). It should be noted that the cell shrinkage and blebbing observed in our study suggests the presence of an active process that has been shown to demand large amounts of ATP in other animal cells where it has been studied (Mills et al., 1999).
In summary, loss of m was detected as the first apoptotic event and was contemporary with early alterations in lysosomic vacuoles. Nuclear and cytoplasmic changes occurred after loss of m and included AVD, exposure of PS on the outer leaflet of the cell membrane, chromatin condensation, rippled nuclear contour, and nuclear segmentation. AVD progressed to blebbing, cell fragmentation, and formation of apoptotic bodies. Later stages were characterized by oligonucleosomal DNA fragmentation. Thus the overall sequence of biochemical events and morphological changes were consonant with current knowledge of apoptotic phenotype in the cells of other animals. The large lysosomic vacuole may represent an extensive reservoir for membrane resealing that is peculiar to coelomocytes. Notably, spontaneous apoptosis in control non-exposed coelomocytes was not detected by any of the morphological or biochemical methods used. To partly explain such low rates of spontaneous apoptosis, one might speculate that viable coelomocytes engulf and rapidly degrade any small number of coelomocytes that undergo apoptosis spontaneously. Sipunculans are oxiconformant worms that have low levels of antioxidant enzymes compared to vertebrates, but also compared to other marine invertebrates, and they are thought to be protected from the formation of reactive oxygen species in mitochondria by a low respiration rate and an alternative, branched electron pathway (Buchner et al., 2001). We have shown that one consequence of overcoming this protective mechanism in coelomocytes by exposure to exogenous H2O2 is the induction of an apoptotic-like mode of cell death.
A normal and well-regulated apoptosis may be essential to an effective innate immune response and the successful elimination of pathogens. In nature, animals can be exposed to a comprehensive range of environmental factors that can result in abnormalities in the regulation of apoptosis, thus compromising the innate immune system (Blanco and Cooper, 2004). Our initial characterization of apoptosis in the sipunculan Themiste petricola may contribute to the future assessment of potential disruptors of apoptosis regulation in this small but geographically widespread phylum of marine worms.
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Acknowledgments
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This work was supported by University of Buenos Aires Grant B081-UBACYT to S.H. and CONICET Grant PEI 6377 to G.B.
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Footnotes
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Received 11 July 2005; accepted 26 October 2005.
¶ Members of the National Research Career (CONICET). 
# Fellow from UBA.
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Abstract
Introduction
