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Vol. 20, Issue 5, 1408-1418, March 1, 2009
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*Department of Biochemistry, Emory University School of Medicine, Atlanta, GA 30322; The Cardiovascular Biology Research Program, Oklahoma Medical Research Foundation, Oklahoma City, OK 73104; and Departmento de Análises Clínicas, Toxicológicas e Bromatológicas da Faculdade de Ciências Farmacêuticas de Ribeirão Preto, University of Sao Paulo, Ribeirão Preto-SP, Brazil
Submitted July 31, 2008;
Revised November 13, 2008;
Accepted December 22, 2008
Monitoring Editor: Donald D. Newmeyer
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Significant expansion of leukocytes involved in both innate and adaptive immunity ultimately results in neutralization and removal of invading pathogens (Nathan, 2006). However, for effective immunological homeostasis to be maintained, efficient contraction of activated leukocyte populations must occur (Antia et al., 2005). Failure to appropriately eliminate activated leukocytes not only enhances the probability of cellular transformation, but also results in leukocyte-mediated damage of viable tissue and can eventually result in autoimmunity (Strasser et al., 2000; Danial and Korsmeyer, 2004).
Many factors regulate leukocyte turnover, including members of the tumor necrosis factor (TNF) and galectin families (Liu and Rabinovich, 2005; Toscano et al., 2007). TNF family members, including Fas, TRAIL, and TNF-
effect leukocyte contraction through the induction of apoptotic cell death. Similarly, several galectin family members, including galectin-3 and -9 (Fukumori et al., 2003; Zhu et al., 2005; Gal-3 and -9), also induce leukocyte removal through apoptosis (Strasser, 2005; Zhu et al., 2005; Stowell et al., 2008c). Cells undergoing apoptotic cell death typically express phosphatidylserine (PS), a phospholipid normally confined to the inner leaflet of the plasma membrane, which serves as a ligand for receptor-mediated phagocytosis (Fadok et al., 2000). Apoptotic cell death also results in DNA degradation and eventual cellular fragmentation (Jacobson et al., 1997; Stroh and Schulze-Osthoff, 1998). Importantly, apoptosis occurs in a coordinated manner, ultimately resulting in homeostatic cellular removal without inciting the deleterious consequences of an inflammatory response (Jacobson et al., 1997).
In contrast to Gal-3, Gal-9, and members of the TNF family, several studies suggest that galectin-1 (Gal-1) may induce PS exposure in leukocytes independent of apoptosis. For example, Gal-1 induces PS exposure in leukocytes in the absence of detectable DNA fragmentation (Dias-Baruffi et al., 2003). Externalization of PS exposure in viable cells may not be limited to Gal-1, as activation of some leukocytes, in particular T-cells and mast cells, can also result in apoptosis-independent PS exposure (Frasch et al., 2004; Elliott et al., 2005; Fischer et al., 2006; Smrz et al., 2007, 2008). Activation-induced PS exposure in viable cells appears to regulate diverse cellular processes ranging from cellular trafficking to proper formation of the immunological synapse (Elliott et al., 2005; Fischer et al., 2006). Taken together, these previous studies strongly suggest that cellular pathways exist whereby PS exposure may be induced in viable cells with diverse consequences.
Although Gal-1–induced PS exposure may occur in viable cells and therefore alter cellular processes as described previously (Frasch et al., 2004; Elliott et al., 2005; Fischer et al., 2006; Smrz et al., 2007, 2008), Gal-1–induced PS exposure sensitizes cells to phagocytic removal (Dias-Baruffi et al., 2003), in a manner similar to cells undergoing apoptotic cell death (Fadok et al., 2001). Such results suggest that Gal-1–induced PS exposure may accompany apoptotic cell death, consistent with several other studies suggesting that Gal-1 actually induces apoptotic cell death in leukocytes (Perillo et al., 1995; Pace et al., 2003). However, Gal-1 is uniquely sensitive to oxidative inactivation. As a result, studies showing Gal-1–induced apoptosis included dithiothreitol (DTT) in cell treatments (Perillo et al., 1997; Pace et al., 2003; Toscano et al., 2007). Indeed, utilization of DTT appears to render cells susceptible to Gal-1–induced cell death (Carlow et al., 2003; Stowell et al., 2007, 2008c). However, the treatment of living cells with DTT can induce the unfolded protein response and can directly induce apoptotic cell death (Braakman et al., 1992; Tartier et al., 2000; Murray et al., 2004), making interpretations about the effects of Gal-1 on leukocyte viability in the presence of DTT difficult.
Although the inclusion of DTT may complicate assays on cellular viability, the ability of Gal-1 to induce PS externalization and prime cells for phagocytic removal, both of which represent characteristic features of cells undergoing apoptotic cell death (Fadok et al., 2000; Dias-Baruffi et al., 2003), suggests that Gal-1 could induce apoptotic cell death, even in the absence of DTT. To help resolve these conflicting interpretations, we have more thoroughly examined whether Gal-1–induced PS exposure occurs in the presence or absence of cell death. In this study we demonstrate that Gal-1–induced PS exposure occurs in the absence of cell death, which suggest a new paradigm in cellular removal.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Gal-1 was alkylated with iodoacetamide followed by removal of free iodoacetamide using a PD-10 column (Amersham Pharmacia Biotech, Piscataway, NJ) equilibrated in PBS, as outlined previously (Stowell et al., 2008a). Gal-1 was labeled using C5 maleimide or carboxylic acid, succinimidyl ester, and dilithium salt reactive dyes (Molecular Probes, Eugene, OR). Before derivatization, β-mercaptoethanol and lactose were removed from Gal-1 using a PD-10 column equilibrated in PBS, followed by incubation with the Alexa-488 reactive dye with 100 mM lactose for 1 h at RT. Free dye was removed by subjection of the reaction mixture to PD-10 column gel filtration.
Cell Culture and Phagocytosis Assay
The isolation of neutrophils was in accordance with a protocol approved by the Emory Institutional Review Board and was accomplished as outlined previously (Stowell et al., 2007). HL60 cells were obtained from ATCC (Manassas, VA) and maintained as outlined previously (Dias-Baruffi et al., 2003). After treatment with the indicated concentrations of galectin, anti-Fas, DTT, lactose, or camptothecin (CAMP), cells were incubated in 50 mM lactose to disengage galectin-treated cells. Cells were labeled with carboxyfluorescein succinimidyl ester (CFSE) as outlined previously (Kaech and Ahmed, 2001). Cell phagocytosis was measured as described previous (Dias-Baruffi et al., 2003).
Flow Cytometric Analysis
For annexin-V (An-V) staining after disengagement, cells were stained in a 100 µl final volume in HEPES buffer (10 mM HEPES, 140 mM NaCl, 5 mM CaCl2, pH 7.4) with a mixture of FITC-conjugated An-V (An-V-FITC; Roche Applied Science, Indianapolis, IN; 2 µl of conjugate per 100 µl of resuspended cells, as described by the manufacturer) and propidium iodide (PI; Molecular Probes; 1 µg/ml final concentration) at 4°C for 15 min. For DNA fragmentation, cells were assessed using the TUNEL reaction (In Situ Cell Death Detection Kit, Roche Applied Science) or hypodiploid analysis as outlined previously (Stowell et al., 2007). Analysis of mitochondrial potential loss using rhodamine-123 was also accomplished as outlined previously (Koya et al., 2000). All samples were assayed using a FACSCalibur flow cytometer (BD Biosciences, San Jose, CA) with a minimum of 10,000 counts/sample. Data were analyzed using Cell Quest software (BD Biosciences).
Cell Sorting
Cells were stained with An-V-FITC, passed through a 70-µM mesh filter to remove any cell aggregates, mounted in a water-cooled jacket to maintain a temperature of 4°C, and analyzed utilizing a MoStar cell sorter (MoFlo, Dako, Denmark, and FacStar Plus, BD Biosciences, at the University of Oklahoma Health Sciences Center). Cells were then physically sorted into An-V–positive and –negative populations based on gates obtained from untreated stained cells. Cells were then washed in Ca2+-free buffer to remove An-V-FITC, examined by flow cytometry, and shown to display no An-V positivity regardless of fraction. Cells were then either restained with An-V-FITC and examined by flow cytometry to determine the integrity of the sorting process or resuspended in complete RMPI and allowed to incubate for the time indicated.
Western Blot Analysis
Cells were treated as indicated followed by SDS-PAGE and transfer to an Immobilon-P membrane (Millipore, Bedford, MA) using a semidry Transblot apparatus (Bio-Rad, Hercules, CA). After transfer, poly(ADP-ribose) polymerase (PARP) was detected using anti-PARP mAb (Novus Biologicals, Littleton, CO).
Confocal Analysis
Cells were incubated with An-V-biotin as outlined above for 1 h at 4°C. After washing, cells were incubated with streptavidin Alexa-488 (Molecular Probes) for 1 h at 4°C. Cells were plated on coverslips pretreated with poly-L-Lysine (Sigma, St. Louis, MO) and allowed to adhere for 30 min at 4°C. Cells were fixed with 2% paraformaldehyde buffered in PBS at 4°C for 2 h as outlined previously (Stowell et al., 2008c). Cells were analyzed using a Leica TCS NT confocal microscope and Leica TCS software (Deerfield, IL). Brightness and contrast adjustment was applied equally to the entire image.
Scanning Electronic Micrograph Analysis
For scanning electronic micrograph (SEM) analysis, neutrophils were treated with Gal-1, PBS, or anti-Fas for 8 h followed by removal of galectin in lactose and fixation in 2% glutaraldehyde in Hanks' balanced salt solution. After fixation, cells were dehydrated by sequential incubation with increasing concentrations of methanol solutions and then dried in liquid carbon dioxide by using a critical point dryer (Autosamdri 814). These fixed dehydrated cells were then mounted on bulk specimen holders (JEOL, Peabody, MA), coated with 60:40 gold/palladium utilizing a Hummer VI sputter coater, and viewed with a JEOL JSM880 SEM under 15-kV accelerating voltage (Oklahoma University, Norman, OK).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Hoffmann et al., 2001). Consistent with previous results, Gal-1 induced robust PS exposure in leukocytes (Figure 1, A and B) as measured by An-V staining after prolonged incubation, similar to cells undergoing apoptotic cell death. However, Gal-1–treated cells displayed complete reversion of PS exposure after Gal-1 removal (Figure 1, A and B), with a cell surface half-life of 
10 h (Figure 1, D–F). By contrast, cells induced to undergo apoptosis by Camp treatment displayed near complete conversion from a PS-positive/PI-negative state to a PS-positive/PI-positive state over the same period of time (Figure 1, A, C, and F).
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). Although neutrophils do not normally phagocytose other neutrophils undergoing apoptotic cell death (Nathan, 2006), we considered the possibility that Gal-1–induced PS exposure may potentially sensitize cells for phagocytic removal by PS-negative cells, similar to recent reports showing that transformed epithelial cells can cannibalize neighboring cells (Overholtzer et al., 2007). In addition, because previous studies suggested that Gal-1 may be mitogenic in some cell populations (Levi and Teichberg, 1983; Adams et al., 1996), we determined whether PS reversion might reflect outgrowth of PS-negative cells. To differentiate between these possibilities, we labeled cells with the intracellular dye CFSE to enable tracking of two distinct cell populations within a mixture. To ensure that CFSE did not alter cellular sensitivity to Gal-1–induced PS exposure, cells were labeled with or without CFSE, followed by incubation with Gal-1. Importantly, CFSE did not alter cellular sensitivity to Gal-1 (Figure 2, A and B), and CFSE-labeled cells could be easily distinguished after Gal-1 treatment within a mixture of cells (Figure 2C). Gal-1–treated PS-positive cells experienced full reversion with no alteration in the percent of cells within each population (Figure 2, D–F). This demonstrates that there was no removal or loss of cells (Barber et al., 2003). In addition, the Gal-1–treated cell population, compared with nontreated cells, displayed nearly identical CFSE dilution (Figure 3, A–C), which is a mark of cell proliferation (Kaech and Ahmed, 2001). These results demonstrate that PS reversion does not represent overgrowth or engulfment by a subpopulation of PS-negative cells and that cells undergoing reversion of PS experience normal cell growth kinetics. Furthermore, physically sorted PS-positive cells after initial Gal-1 treatment reverted PS upon removal of Gal-1, similar to cells mixed with untreated cells (Figure 3, A–E). This reversion occurred without alterations in DNA degradation or cell growth (Figure 3, D, F, and G), which further demonstrated that Gal-1–induced PS exposure autonomously reverts after Gal-1 removal.
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). Consistent with this, Gal-1 failed to alter mitochondrial potential, although cells undergoing apoptotic cell death demonstrated significant depolarization (Figure 5, A and C). Furthermore, treatment of cells with the pan-caspase inhibitor, zVAD-fmk, did not inhibit Gal-1–induced PS exposure (Figure 5, B and D), although it inhibited PS exposure in cells undergoing apoptotic cell death (data not shown). In addition, cells treated with Gal-1 failed to display cleavage of the common caspase substrate PARP (Lazebnik et al., 1994); by contrast, cells undergoing apoptotic cell death displayed significant PARP cleavage (Figure 5E). These results show that Gal-1 induces PS exposure independent of irreversible processes, such as mitochondrial potential loss or caspase activation, consistent with the ability of PS to revert in its surface exposure after Gal-1 treatment and withdrawal and strongly suggesting that Gal-1–induced PS exposure occurs independent of cell death.
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). Previous results suggested that Gal-1–induced PS exposure may exhibit some asymmetric distribution on the plasma membrane, similar to early phases of apoptotic cell death (Godard et al., 1999), suggesting that cells incubated with Gal-1 may fail to transition to later phases of apoptosis where greater PS positivity occurs. To examine this in more detail, we analyzed cells by confocal analysis for localization of PS expression after incubation with Gal-1 at later time points. In contrast to cells undergoing apoptotic cell death, Gal-1–treated cells retained punctate PS exposure, whereas cells undergoing apoptosis progressed to nearly uniform PS exposure (Figure 6A).
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). We examined whether cell surface alterations that normally accompany apoptosis might also occur during Gal-1–induced PS exposure. Consistent with previous results (Scott et al., 2001), SEM analysis of apoptotic cells showed significant retraction of microvilli and cell surface flattening with prominent formation of apoptotic blebs (Figure 6, D, G, and H). By contrast, cells exposed to Gal-1 displayed morphological features indistinguishable from nonapoptotic controls (Figure 6, B, C, E, and F). These results support the conclusion that Gal-1 induces PS exposure independent of cell death and without changes in cellular morphology.
Although PS exposure induced by Gal-1 clearly reverts after Gal-1 removal, we sought to define whether continuous Gal-1 incubation could induce sustained PS exposure over prolonged incubation periods. However, Gal-1 loses significant activity after prolonged incubation periods (days; Hirabayashi and Kasai, 1991; Cho and Cummings, 1995a). Previous studies utilized DTT in cell treatments to prevent Gal-1 oxidation (Perillo et al., 1995; Pace et al., 2003; Toscano et al., 2007). However, DTT alone can induce significant alterations in cellular viability and artificially sensitize cells to Gal-1–induced apoptosis (Figure 7, A and B; Tartier et al., 2000; Stowell et al., 2007, 2008c). As a result, we stabilized Gal-1 by derivatization with iodoacetamide (iGal-1), as outlined previously (Whitney et al., 1986; Stowell et al., 2008c). Similar to the unmodified protein, iGal-1 induced significant PS exposure in cells (Figure 7C). iGal-1–induced PS exposure required binding to cell surface carbohydrates, because lactose, a general inhibitor of galectin-ligand interactions (Stowell et al., 2008b), completely blocked iGal-1–induced PS exposure (Figure 7C). Furthermore, iGal-1–induced PS exposure displayed equivalent kinetics, occurred over a similar dose response, and also sensitized cells to phagocytosis at levels comparable to cells undergoing apoptotic cell death as observed previously (Figure 7, D–F; Dias-Baruffi et al., 2003).
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; Stowell et al., 2007, 2008c). Cells incubated with iGal-1 in the absence of DTT maintained continual PS exposure over 3 d (Figure 8, A and D). Importantly, although iGal-1–induced sustained PS exposure, no detectable changes in PI staining, cellular fragmentation, or DNA degradation occurred (Figure 8, A–F). By contrast, cells undergoing apoptosis displayed significant increases in PI staining, cellular fragmentation, and DNA degradation over the same time period (Figure 8, A–F).
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). Cells treated with iGal-1 continued to experience CFSE dilution while maintaining uniform PS exposure over time (Figure 9, A and C). PS exposure also remained punctate over 2 d of continuous iGal-1 incubation (Figure 9B) with no alterations in cell cycle progression or viable cell number (Figure 9, D and E). Taken together, these results demonstrate that iGal-1 induces sustained PS exposure without altering cell growth or viability.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Figure 7, D–F), these results demonstrate that Gal-1 induces phagocytic removal of living cells, as opposed to apoptotic cells, suggesting a new paradigm in cellular turnover.
Although many studies suggested that Gal-1 affects leukocyte turnover and function (van Kooyk and Rabinovich, 2008), whether Gal-1 directly alters leukocyte viability remained enigmatic. Many previous studies utilized the reducing agent DTT when evaluating the effect of Gal-1 on leukocyte viability (Perillo et al., 1995; Perillo et al., 1997; Pace et al., 2003; Toscano et al., 2007). However, DTT induces the unfolded protein response, directly induces apoptosis, and sensitizes cells to Gal-1–induced cell death (Braakman et al., 1992; Tartier et al., 2000; Murray et al., 2004; Stowell et al., 2007, 2008c). Because the extracellular environment is largely oxidative and DTT artificially penetrates cell membranes (Kemp et al., 2008), the physiological relevance of DTT inclusion remains unclear.
Other studies demonstrate that Gal-1 induces PS exposure and sensitizes cells to phagocytic removal in the absence of DTT (Dias-Baruffi et al., 2003), suggesting that Gal-1 may alter cell viability regardless of DTT inclusion. However, these studies failed to detect DNA fragmentation or alterations in membrane integrity after Gal-1 incubation (Dias-Baruffi et al., 2003; Stowell et al., 2007, 2008c), suggesting that either definitive signs of Gal-1–induced cell death required prolonged Gal-1 incubation or that Gal-1 induces phagocytic removal of living cells. To clarify the effect of Gal-1 on cell viability we utilized iGal-1, which retains previously defined activities of nonderivatized Gal-1, yet remains resistant to oxidation in the absence of DTT (Whitney et al., 1986; Dias-Baruffi et al., 2003; He et al., 2003; Stowell et al., 2008c). Cells treated with iGal-1 maintained PS exposure for 3 continuous days without inducing changes in membrane integrity, DNA degradation, or cell size. Furthermore, cells continued to growth normally as indicated by absolute viable cell numbers, CFSE dilution, and cell cycle analysis. These results not only demonstrate that Gal-1 fails to alter cell viability while maintaining prolonged PS externalization, but also suggest that PS asymmetry is not a strict requirement for cell division.
The ability of Gal-1 to induce PS exposure despite failing to alter cellular viability presented the unusual possibility that cells incubated with Gal-1 may reverse PS exposure after Gal-1 removal. This was important to definitively evaluate, as PS externalization is commonly thought to be a terminal event of, and directly correlated with, cells undergoing apoptotic cell death (Fadok et al., 2000; Hoffmann et al., 2001). Indeed, removal of Gal-1 resulted in PS reversion without detectable alterations in cell viability, which further demonstrated the nonapoptotic nature of Gal-1–induced PS exposure. Importantly, although several studies suggest that Gal-1 may be mitogenic at low concentrations (Levi and Teichberg, 1983; Adams et al., 1996), reversion did not represent mitogenic outgrowth of a PS negative population, as differentially labeled cells and physically sorted cells autonomously reverted PS. The ability of PS-positive cells to autonomously revert PS also ruled out the possibility that PS reversion reflected cannibalism by bystander cells, as recently reported to occur in transformed epithelial cells (Overholtzer et al., 2007). The ability of Gal-1 to induce PS exposure despite failing to engage irreversible pathways commonly associated with apoptotic cell death, such as caspase activation and loss of mitochondrial potential (Hengartner, 2000), also corroborates the reversible nature of Gal-1–induced PS exposure. Equally important, although some studies demonstrate that caspases may directly induce PS exposure (Martin et al., 1996; Vanags et al., 1996), caspase activation is not a prerequisite or necessary for PS exposure to occur.
In addition to inducing PS exposure in the absence of apoptosis, Gal-1–induced PS exposure morphologically differed from PS exposure accompanying cell death. Gal-1–induced PS exposure remained punctate regardless of the duration of Gal-1 incubation, whereas cells undergoing apoptotic cell death displayed uniform PS exposure before transition to late apoptosis and loss of membrane integrity. Previous results demonstrate that apoptotic cells also exhibit punctate PS exposure during early phases of apoptosis, with gradual progression over the entire cell surface with time (Godard et al., 1999). Although the exact players responsible for PS mobilization remain uncertain (Schlegel and Williamson, 2007), PS asymmetry does require an ATP driven aminophospholipid translocase (APT) activity that translocates any external PS back to the inner leaflet of the plasma membrane (Dolis et al., 1997). By contrast, putative phospholipid scramblases, which are thought to facilitate PS externalization, often occur in membrane microdomains (Damek-Poprawa et al., 2006). Failure of cells incubated with Gal-1 to undergo cell death may prevent full inactivation of ATP-driven APT (Seigneuret and Devaux, 1984; Gleiss et al., 2002), suggesting that Gal-1 may activate PS exposure in areas near lipid rafts, followed by reinternalization by incompletely inactivated APT, resulting in the appearance of punctate PS. Although Gal-1–induced PS exposure occurs independent of caspase activation, cells undergoing apoptosis activate putative scramblases through caspase-mediated activation of protein kinase C (PKC) that results in scramblase phosphorylation and activation (Frasch et al., 2000). The ability of Gal-1 to activate PKC (Karmakar et al., 2005) suggests that Gal-1 may be able to induce PS exposure independent of caspase activation by directly activating scramblase.
Gal-1 displays unique sensitivity to oxidative inactivation. Thus, increases in environmental oxidation associated with failure to remove pathogens may trigger reversal of PS exposure in leukocytes originally targeted for removal to allow reengagement in host defense (Hirabayashi and Kasai, 1991; Cho and Cummings, 1995b; Nathan, 2006; Kemp et al., 2008). Such normally oxidative environments may be especially important for granulocytes, which do not possess antigen specific immunity and therefore have limited capacity to directly detect pathogen levels (Nathan, 2006). Failure to remove pathogens results in tissue damage and injury, which causes additional leukocyte recruitment and enhanced oxidation of the extracellular environment. Enhanced oxidation leads to Gal-1 inactivation (Inagaki et al., 2000), allowing cells originally targeted for removal to revert PS and become reengaged in host defense. Consistent with this, continual incubation of cells with nonstabilized Gal-1 results in spontaneous Gal-1 oxidation and PS reversion (Stowell and Cummings, unpublished observations). Thus, by such mechanisms Gal-1 may be uniquely suited to sense oxidative environments and alter leukocyte turnover and accumulation.
Although apoptosis generally prevents inflammation during cellular turnover, exuberant apoptosis, as can occur during inflammation, can actually be proinflammatory (Misawa et al., 2001). Consistent with this, several studies suggest that granulocytes have a unique apoptosis-independent pathway(s) of removal. Granulocytes transgenically overexpressing bcl-2 fail to undergo apoptotic cell death, yet display normal sensitivity to phagocytic removal in vivo and in vitro (Lagasse and Weissman, 1994). Furthermore, Kupffer cells phagocytose nonapoptotic neutrophils in vivo (Shi et al., 2001). In addition, Fas and FasL null mice, which display significant defects in lymphocyte turnover, display no alterations in homeostatic granulocyte turnover, both in resting and inflammatory conditions (Fecho and Cohen, 1998). In contrast to the reversible nature of Gal-1–induced PS exposure, the ability of Gal-1 to sustain PS exposure may reflect successful pathogen removal, inflammation resolution, and therefore reduced Gal-1 oxidation. In this way, Gal-1–induced turnover of viable cells allows granulocytes to actively maintain membrane integrity until successfully phagocytosed.
In addition to regulating leukocyte turnover, recent studies suggest that Gal-1–induced PS exposure in viable cells may also regulate other fundamental cellular processes. For example, PS exposure during T-cell activation appears to alter ion transport, cellular trafficking, and immunological synapse formation (Elliott et al., 2005; Fischer et al., 2006), although the extent to which PS directly modulates these processes remains uncertain (Martinez et al., 2006). In addition to T-cells, several studies suggest that B cells may also externalize PS independent of apoptosis (Hammill et al., 1999; Dillon et al., 2000, 2001; Elliott et al., 2006). Brief PS externalization may also facilitate the release of preformed inflammatory mediators, as macrophages and mast cells display transient PS exposure after activation (Martin et al., 2000; MacKenzie et al., 2001). By contrast, recent studies demonstrated that ligation of cell surface GPI (glycosylphosphotidylinositol)-anchored proteins on mast cells can induce sustained PS exposure independent of cell death (Smrz et al., 2007, 2008). Similar to Gal-1–induced PS exposure, sustained PS exposure in mast cells may contribute to the removal of viable cells harboring pathogens during infection (Smrz et al., 2007).
Before the discovery of alterations in PS asymmetry in cells undergoing apoptosis (Schlegel and Williamson, 2001), many studies documented a key role for PS asymmetry on activated platelets in thrombosis (Monroe et al., 2002). Similar to platelets, PS exposure on cells can also induce thrombosis (Tracy et al., 1985; Williamson et al., 1992), which suggests that in addition to regulating neutrophil turnover, Gal-1–induced PS exposure may also contribute to fibrin deposition. Consistent with this, improper resolution of inflammatory responses often results in significant peripheral fibrin deposition (Loeuille et al., 2005), suggesting a role for PS-positive neutrophils awaiting removal in this process.
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ACKNOWLEDGMENTS |
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Footnotes |
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Address correspondence to: Richard D. Cummings (rdcummi{at}emory.edu)
Abbreviations used: An-V, annexin V; Camp, camptothecin; CFSE, carboxyfluorescein diacetate, succinimidyl ester; DTT, dithiothreitol; Gal-1, galectin-1; Gal-3, galectin-3; Gal-9, galectin-9; PARP, poly(ADP-ribose) polymerase; PI, propidium iodide; PS, phosphatidylserine; SEM, scanning electron microscopy; TNF, tumor necrosis factor.
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