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The Journal of Immunology, 2005, 174: 3999-4009.
Copyright © 2005 by The American Association of Immunologists

Effector CD4+ T Cells Generate Intermediate Caspase Activity and Cleavage of Caspase-8 Substrates1

Ravi S. Misra*, Dawn M. Jelley-Gibbs{dagger}, Jennifer Q. Russell*, Gail Huston{dagger}, Susan L. Swain{dagger} and Ralph C. Budd2,*

* University of Vermont, Immunobiology Program, College of Medicine, Burlington, VT 05405; and {dagger} Trudeau Institute, Saranac Lake, NY 12983


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Caspase-8 activation promotes cell apoptosis but is also essential for T cell activation. The extent of caspase activation and substrate cleavage in these divergent processes remains unclear. We show that murine effector CD4+ T cells generated levels of caspase activity intermediate between unstimulated T cells and apoptotic populations. Both caspase-8 and caspase-3 were partially activated in effector T cells, which was reflected in cleavage of the caspase-8 substrates, c-FLIPL, receptor interacting protein 1, and to a lesser extent Bid, but not the caspase-3 substrate inhibitor of caspase-activated DNase. Th2 effector CD4+ T cells manifested more caspase activity than did Th1 effectors, and caspase blockade greatly decreased initiation of cell cycling. The current findings define the level of caspase activity and substrates during initiation of T cell cycling.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
It is becoming increasingly evident that ligation of death receptors and activation of cellular caspases does not necessarily always result in apoptosis. Involvement of the death receptor, Fas, in processes other than cell death has been revealed by the ability of Fas ligation to enhance neural outgrowth (1), cardiac hypertrophy (2), liver regeneration (3), fibroblast proliferation (4), dendritic cell maturation (5), and T cell activation (6). More directly, caspase-8 plays an integral role in T cell function, as humans bearing nonfunctional caspase-8, as well as caspase-8 conditional knockout mice, exhibit severe defects in T cell activation (7, 8). However, there is currently no clear picture of which caspases or which substrates are involved with T cell activation and how far down the caspase cascade these signals propagate.

Traditionally, activation of the upstream initiator caspases that possess large prodomains, such as procaspases-8, -9, and -10, had been thought to require proteolytic cleavage of the zymogen, resulting in the release of large and small subunits (9). These fragments then heterodimerize to form an active caspase molecule. Typically, the cleavage occurs by auto- and cross-proteolysis in response to the ligation of death receptors, which increases the local concentrations of caspases (9, 10, 11). The upstream initiator caspase signal is propagated to the effector procaspases, such as procaspase-3, by proteolytic cleavage of the procaspase. As part of this caspase cascade, various caspase substrates are cleaved, including the DNase inhibitor inhibitor of caspase-activated DNase (ICAD)3 (12) and structural proteins, such as actin (13), fodrin (14), and lamin (15).

The earlier view that caspase activation occurs only through its cleavage has been revised for certain upstream caspases. An alternative method of caspase activation is through a conformational change. For example, when caspase-9 interacts with cytochrome c complexed to Apaf-1, a conformational change of caspase-9 exposes its active site (16, 17). In a similar manner, procaspase-8 can induce a structural change in a neighboring procaspase-8 molecule, resulting in active full-length caspase-8 (10). In addition, the enzymatically inactive casapase-8-like molecule c-FLIPL can also associate with and activate full-length caspase-8 (18). As c-FLIPL also contains a known caspase cleavage site, it represents potentially one of the earliest substrates of caspase-8 (19).

The significance of caspase activity in nonapoptotic effector T cells is still poorly understood. It is not known how the levels of active caspases or cleavage of their substrates in viable effector T cells compare with those in resting T cells or those undergoing apoptosis after Fas ligation. We observe that intermediate levels of caspase activity are generated at the effector stage of T cell activation, which are proportional to cell cycling, and that blockade of caspase activity inhibits cell cycling. Proximal caspase-8 substrates such as c-FLIPL and receptor interacting protein 1 (RIP1) are cleaved in effector T cells, but not the downstream caspase-3 substrate ICAD. Furthermore, the caspase activity and cleavage of substrates is more extensive in Th2 effector cells than in Th1 effectors. Collectively, these results define the caspase activity profile in viable T cells and underscore the functional requirements of such activity.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice

C57BL/6 mice were housed and bred in the University of Vermont animal facility and were used at 2–6 mo of age. B10.Br V{alpha}11/V{beta}3 AND TCR transgenic (pigeon cytochrome c/I-Ek responsive) mice were used at 2–3 mo of age and were bred in the animal facilities at the Trudeau Institute. Both facilities are American Association of Laboratory Animal Care approved and protocols were approved by the Institutional Animal Care and Use Committee at the respective institutions.

T cell purification

C57BL/6 spleens and lymph nodes were isolated and disrupted through nylon mesh in RPMI 1640 (MediaTech) containing 5% (v/v) FCS (HyClone). Erythrocyte lysis of splenocytes was performed using Gey’s solution. Lymphocytes and splenocytes were combined and CD4+ T cells were isolated by negative selection. C57BL/6 cells were incubated with anti-CD8 (Tib 105), anti-MHC class II (M5/114/15/2; a kind gift of M. Rincón, University of Vermont, Burlington, VT), anti-CD11b (M1/70), anti-NK1.1 (PK136), and anti-B220 (RA3-6B2) on ice for 45 min. For total T cell isolation, anti-CD8 was omitted from Ab mix. Cells were washed three times and rocked with goat anti-mouse and goat anti-rat conjugated magnetic beads at a 10:1 ratio of beads to cell (Qiagen) at 4°C for 45 min. Magnetic depletion was used to remove bead-bound cells, routinely yielding >90% CD4+ T cells. Cells were washed and resuspended in culture medium (RPMI 1640 supplemented with 25 mM HEPES, 2.5 mg/ml glucose (Sigma-Aldrich), 10 µg/ml folate (Invitrogen Life Technologies), 110 µg/ml pyruvate (Invitrogen Life Technologies), 5 x 10–5 M 2-ME (Sigma-Aldrich), 292.3 µg/ml glutamine (Invitrogen Life Technologies), 100 U/ml penicillin-streptomycin (Invitrogen Life Technologies), and 5% FBS).

Naive AND CD4+ T cells were enriched from pooled spleen and lymph node cells as previously described (20). In brief, spleen and lymph node cells were passed through a nylon wool column, and nonadherent cells were treated with a panel of CD8, HSA, and MHC class II depleting Ab and complement followed by Percoll (Amersham Biosciences) centrifugation. CD4+ purity was routinely >95%, 90–95% of which had a naive phenotype (CD45RBhigh, CD62 ligandhigh, CD44low, CD25low) and expressed the TCR transgene. AND transgenic cells were cultured in RPMI 1640 plus penicillin (200 µg/ml; Sigma-Aldrich), streptomycin (200 µg/ml; Sigma-Aldrich), glutamine (4 mM; Sigma-Aldrich), 2-ME (50 µM; Sigma-Aldrich), HEPES (10 mM; Sigma-Aldrich), and 8% FBS (Intergen).

T cell culture

C57BL/6 CD4+ T cells were activated in culture medium by plate-bound anti-CD3 (5 µg/ml; 145-2C11), anti-CD28 ascites (1:500), and recombinant human IL-2 (50 U/ml; Cetus) for 2 days. Cells were then removed from anti-CD3 and fed with fresh medium plus IL-2. AND effectors were generated by culturing transgenic CD4+ cells as previously described (20). Alternatively, AND CD4+ T cells were stimulated with DCEK-ICAM APCs (1.5 x 105 cells/ml) pulsed with 5 µM pigeon cytochrome c fragment (PCCF; KAERADLIAYLKQATAK) and IL-2 (11 ng/ml). The DCEK-ICAM cell line was originally generated by R. Germain (National Institutes of Health, Bethesda, MD) and expresses B7.1, ICAM-1, and class II MHC (I-Ek). APCs were treated with 100 µg/ml mitomycin C (Sigma-Aldrich) for 45 min at 37°C, extensively washed, and then used at a 2:1 T cell/APC ratio.

Murine IL-2 and IL-4 were obtained from culture supernatant of X63.Ag8-653 cells transfected with cDNA for the respective cytokines. Th1 culture conditions included 10 µg/ml anti-IL-4 neutralizing Ab (11b11; BD Biosciences) and 4 ng/ml IL-12 (R&D Systems). Th2 cells were stimulated in the presence of 10 µg/ml anti-IFN-{gamma} neutralizing Ab (XMG1.2; BD Biosciences) and 30 ng/ml IL-4 (R&D Systems). Recombinant murine IL-12 used in AND T cell experiments was a gift of S. Wolf (Genetics Institute, Cambridge, MA). Where indicated, culture conditions included the pan-caspase blocker z-Val-Ala-Asp-fluoromethylketone (z-VAD; Enzyme System Products; 100 µM) or comparable DMSO (Sigma-Aldrich) vehicle control. Viable cells were isolated by layering cell suspensions over Lympholyte-M (Cedarlane Laboratories) and centrifugation at 850 x g for 20 min. Cells were washed three times and cell counts performed.

RIP1-deficient 3T3 cell culture

RIP1-deficient 3T3 cells were a kind gift of M. Kelliher (University of Massachusetts, Worcester, MA) and were grown in DMEM supplemented with Hams 3F12 (Invitrogen Life Technologies), 5 x 10–5 M 2-ME (Sigma-Aldrich), 292.3 µg/ml glutamine (Invitrogen Life Technologies), 100 U/ml penicillin-streptomycin (Invitrogen Life Technologies), and 10% FBS (HyClone).

Caspase activity assays

Relative caspase activities were determined per vendor recommendation using the Apo-ONE Caspase-3/7 Assay (Promega). Viable cells were isolated and resuspended in complete medium at 10 x 106/ml. Cells were serially diluted in complete medium and then mixed with 100 µl of Caspase-3/7 Reagent (DEVD-rhodamine) per the manufacturer’s protocol. Competition experiments included 100 µM z-VAD in the Caspase-3/7 Reagent mix. Spectrophotomeric readings were taken over a range of times and data shown represent readings at 90 or 150 min.

Immunoblot analysis

Viable cells were lysed in buffer containing 0.2% Nonidet P-40, 20 mM Tris-HCl (pH 7.4), 2 mM sodium orthovanadate, 10% glycerol, 150 mM NaCl, complete protease inhibitor (Roche Diagnostics), and 10 µM z-VAD (Enzyme System Products). Protein concentration was determined by Bradford assay (Bio-Rad). Protein lysates were boiled for 5 min in loading buffer containing 2-ME and were separated using SDS-PAGE on 12.5% gels. Proteins were transferred onto a polyvinylidene difluoride membrane (Bio-Rad) and blocked using 4% milk in TBS plus 0.1% Tween 20 at room temperature for 1 h. Membranes were incubated at 4°C overnight in milk containing one of the following primary detection Abs: 1 µg/ml actin (I-19; Santa Cruz Biotechnology), Bid (R&D Systems), caspase-3 (a kind gift of Y. Lazebnik, Cold Spring Harbor Laboratories, Cold Spring Harbor, NY), caspase-8 (a kind gift of A. Stasser, The Walter and Eliza Hall Institute of Medical Research, Melbourne, Australia), ICAD (BD Biosciences), RIP1 (BD Biosciences), or 2 µg/ml FLIP (Dave-2; Apotech). Blots were washed three times for 15 min each and incubated for 90 min with 0.5 µg/ml (caspase blots) or 1 µg/ml (all others) of appropriate secondary Ab conjugated to HRP (Santa Cruz Biotechnology). Blots were washed again and developed using LumiGLO (Kirkegaard & Perry Laboratories).

Biotin-VAD-fmk caspase precipitation assay

Viable T cells were incubated with z-FA-fmk control peptide or 100 µM nonbiotinylated z-VAD at 37°C for 15 min followed by another incubation with 10 µM biotin-VAD (Enzyme System Products). Cells were lysed in buffer containing 20 µM biotin-VAD. A total of 600 µg of lysate was then precleared by rocking with 40 µl of Sepharose 6B agarose beads (Sigma-Aldrich) at 4°C for 2 h. Supernatant was then rocked with 30 µl of Strepavidin-Sepharose beads (Zymed Laboratories) at 4°C overnight. Beads were washed five times in lysis buffer without protease inhibitor, and then were boiled in loading buffer. Beads were removed by centrifugation, and immunoblot analysis was then performed on supernatants.

CFSE dye labeling of cells

Cells were washed with PBS plus 0.1% BSA and then were mixed with 5 µM CFSE in PBS plus 0.1% BSA (Molecular Probes). Samples were divided into 1-ml aliquots and incubated for 10 min at 37°C. Cells were then washed three times. A portion of cells was incubated overnight at 37°C to serve as a noncycling control for flow cytometry, and the remaining cells were stimulated.

Flow cytometry

For surface staining, cells were washed in PBS and then incubated with 1 µg/ml FITC-conjugated anti-CD4 Ab diluted in PBS/1% BSA for 30 min. Cells were then washed with PBS/BSA and fixed using 1% formaldehyde in PBS. Samples were analyzed using an LSRII flow cytometer (BD Biosciences).

For TUNEL and Bcl-2 staining, surface staining was first completed as above, with the exception that cells were fixed on ice for 15 min using 2% formaldehyde in PBS. Cells were then washed twice with PBS, fixed in 70% ice-cold ethanol for 15 min, and then washed twice with PBS. Nicked DNA was labeled by incubating cells with TdT buffer (2.5 mM CoCl2, 1 U TdT, and 0.5 nmol biotin-dUTP (Roche Diagnostics) in a total volume of 50 µl) at 37°C for 1 h. Cells were then washed twice with 1% BSA in PBS and subsequently incubated with Strepavidin-Tricolor for 30 min. Samples were then washed twice with 1% BSA in PBS and fixed using 1% formaldehyde (Ted Pella) in PBS. For further Bcl-2 staining, cells were permeabilized with 1% BSA and 0.03% saponin (Sigma-Aldrich) in PBS on ice for 10 min. Cells were then washed and incubated with FITC-conjugated anti-Bcl-2 (BD Biosciences) on ice for 30 min. Samples were washed three times and fixed using 2% formaldehyde (Ted Pella) in 1% BSA in PBS. For staining of active caspase-3 intracellular staining, surface staining was first completed and cells were fixed. Cells were washed twice after fixation and the Anti-Active Caspase-3 Apoptosis kit 1 (BD Biosciences) was used to detect intracellular active caspase-3, with PBS/0.03% saponin used in place of Perm/Wash buffer. Cells were washed twice with PBS before TUNEL staining.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Effector CD4+ T cells possess active caspases

Although caspase activity is essential for T cell activation (7, 21, 22), the extent of caspase activation and their substrate cleavage have not been defined. Caspase activation and cleavage of known substrates were examined during the progression of naive CD4+ T cells to the effector stage. Fresh polyclonal CD4+ T cells were purified from C57BL/6 mice, a portion were stimulated with anti-CD3/CD28 in the presence of IL-2, and nonviable cells were removed at all time points by Ficoll centrifugation. Caspase activity was then determined using a DEVD-rhodamine substrate (Fig. 1A). Although DEVD has a high affinity for active caspase-3, it also has an appreciable affinity for the other caspases, including caspase-8 (23). This assay is therefore best viewed as an indication of overall caspase activity (9, 24). Fresh CD4+ T cells had negligible levels of caspase activity, whereas viable day 4 effectors manifested considerably increased caspase activity (Fig. 1A). Caspase activity in effector CD4+ T cells was greatly diminished in the presence of the pan-caspase blocker z-VAD-fmk (Fig. 1B), a competitive inhibitor of DEVD, confirming the specificity of the assay. In contrast, apoptotic T cells generated by incubating T cell blasts with soluble Fas ligand (FasL) manifested a 4-fold increase in caspase activity compared with viable effector T cells (Fig. 1C). Intracellular levels of active caspase-3 were assessed in fresh day 4 effectors, and day 4 effectors were treated with FasL (Fig. 1D). This demonstrated monophasic peaks of active caspase-3, whose mean fluorescence intensity increased from 82 in fresh cells to 825 in day 4 effectors and 1717 in effectors treated with FasL. Cells were simultaneously stained for TUNEL, which revealed 3% of fresh cells, 9% of effectors, and 63% of FasL-treated cells were TUNEL+. This demonstrated that the level of caspase activity in effector CD4+ T cells was intermediate between that of resting T cells and of those undergoing programmed cell death. To determine the levels of active caspase-3 in truly nonapoptotic TUNEL– effectors, TUNEL– and TUNEL+ populations of effector cells were gated separately and caspase-3 activity was measured. Active caspase-3 levels were clearly higher in TUNEL+ cells, but TUNEL– cells had levels of active caspase-3 that were still substantially above those of fresh T cells (Fig. 1E).



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  		FIGURE 1. Intermediate levels of caspase activity are present in effector CD4+ T cells. Purified polyclonal CD4+ T cells were stimulated for 2 days with immobilized anti-CD3, soluble anti-CD28, and IL-2, and then were expanded in fresh medium supplemented with IL-2 for an additional 2 days. A, Viable unstimulated (Fresh) and day 4 effector CD4+ T cells were purified by ficoll centrifugation and caspase activity was determined using a DEVD-rhodamine fluorogenic assay. B, To confirm specificity of the assay, the broad spectrum caspase inhibitor z-VAD-fmk (100 µM) was used as a competitive inhibitor of DEVD-rhodamine using day 4 effector cells. Results are representative of five experiments. C, As a positive control for caspase activation and as a point of reference, a portion of the same effector T cells was treated with FLAG-tagged FasL cross-linked with anti-FLAG Ab for 4 h before the caspase assay was performed. D, Fresh CD4+ T cells (peak 1), day 4 effector cells (peak 2), and 2-h FasL-treated effector cells (peak 3) were simultaneously analyzed for intracellular active caspase-3 and TUNEL. E, The day 4 effector cells were further subdivided into TUNEL– and TUNEL+ subsets, and levels of active caspase-3 were displayed for each. Note that the nonapoptotic TUNEL– effector cells nonetheless have higher levels of active caspase-3 than do fresh cells. Results in D and E are representative of two independent experiments.

 
To further define which caspases contributed to the caspase activity of day 4 effector CD4+ T cells, we examined the activation of upstream caspase-8 and downstream caspase-3 as defined by their cleavage at specific aspartate residues. We also determined cleavage of the known caspase-8 substrates c-FLIPL and Bid as well as the caspase-3 substrate ICAD by immunoblot using lysates from purified CD4+ T cells that were either unstimulated or activated by anti-CD3/CD28 (Fig. 2). Lysates from FasL-treated cells were included as a positive control for substrate cleavage products. One means of caspase-8 activation is by cleavage of its full-length 55-kDa form, resulting in p43/41 caspase-8, through the liberation of a C-terminal p10/12 fragment. Further cleavage of p43/41 caspase-8 leads to the generation of a p18 fragment that heterodimerizes with the p10 fragment to form a fully active protease (25). Caspase-8 was largely present in its full-length 55-kDa form in freshly isolated CD4+ T cells (Fig. 2A). Of note, p18 caspase-8 was absent in effector lysates, but was present in lysates from the FasL-treated population (Fig. 2A). Cleavage of caspase-8 to p43/41 caspase-8 was seen 2 days after activation. Evidence of caspase-8 cleavage was apparent as early as 4 h after T cell activation (21). The intensity of the p43/41 complex increased gradually through day 4 and then decreased slightly by day 6 (Fig. 2A). The kinetics of the cleavage pattern closely paralleled CD25 expression and cell cycle rates (Ref. 21 and data not shown).



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  		FIGURE 2. Caspase activation and substrate cleavage are concomitant with CD4+ T cell activation. Day 4 polyclonal effector CD4+ T cells were generated as in Fig. 1. Lysates were prepared from viable cells at the times indicated. Immunoblot analysis was performed for caspase-8 (A), c-FLIPL (B), Bid (C), caspase-3 (D), and ICAD (E). Caspase-8 represents noncleaved procaspase-8, whereas p43/41caspase-8 bands represent fragments generated by cleavage at Asp374 or Asp384. c-FLIPL represents full-length protein, p43FLIP represents a fragment generated by cleavage at Asp376, and p27FLIP represents a further cleavage product from cleavage at Asp258. t-Bid is generated by cleavage of full-length Bid at Asp75. Caspase-3 represents noncleaved procaspase-3, whereas p20 caspase-3 and p12/10 caspase-3 represent fragments generated by cleavage at Asp169 or Asp175. ICAD represents noncleaved ICAD, and p18ICAD is generated by cleavage at Asp117. Lysates from FasL-treated populations were included in the analysis to illustrate cleavage products. NS, Nonspecific band

 
c-FLIPL heterodimerizes with caspase-8 and is a caspase-8 substrate (19, 26). In a manner similar to caspase-8, cleavage of c-FLIPL from its full-length 55-kDa form to p43FLIP was considerably more apparent by day 2 (Fig. 2B). This corresponds with a known caspase-8 cleavage site in c-FLIPL at Asp376 (27, 28). Bid was only weakly expressed in fresh T cells, but was considerably up-regulated by day 2 after activation (Fig. 2C). Cleavage of Bid was only slight but mirrored that of caspase-8 activation, becoming apparent at day 2, peaking at day 4, followed by a gradual decrease by day 6 (Fig. 2C). Thus, caspase activation in effector CD4+ T cells led to the cleavage of the known caspase-8 substrate, c-FLIPL, and less extensively downstream, Bid.

Caspase-3 is inactive in its full-length 33-kDa form and is cleaved by upstream caspases, including caspase-8, which liberates a large p20 and a small p12/10 fragment (25, 29). The large and small fragments then dimerize to form the active protease (25). p20 caspase-3 may then be autoproteolytically processed to a p17 fragment, which is considered the mature large caspase-3 subunit (29, 30, 31). In fresh CD4+ T cells, full-length caspase-3 and p20 caspase-3 were present, whereas active p12/10 caspase-3 was not apparent until day 2 (Fig. 2D). p17 caspase-3 was never observed in the immunoblot of whole cell lysates, even those treated with FasL. This may be explained by the lack of ability of the anti-caspase-3 Ab to strongly detect p17 caspase-3. However, in lysates made from cells treated with biotin-VAD, a 17-kDa band was revealed with HRP-conjugated Strepavidin or anti-biotin Ab and was enriched after the precipitation of active caspases, which provides support for p17 caspase-3 being catalytically active in effector T cells (see Fig. 3, and data not shown). Despite the degree of caspase-3 cleavage, only minimal cleavage of ICAD was detectable (Fig. 2E). Cleavage of each of these caspase substrates was considerably less than that observed in the same cells treated with FasL (Fig. 2).



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  		FIGURE 3. Activated T cells possess active full-length caspase-8, active cleaved caspase-3, full-length FLIPL, p43FLIP, and RIP1 complexed with active caspases. Freshly isolated polyclonal T cells (lanes 1 and 3) or day 4 effectors (lanes 2, 4, and 5) were incubated with control peptide z-FA-fmk (lanes 1-4) or 100 µM nonbiotinylated z-VAD (lane 5) at 37°C for 15 min. All cells were then incubated with 10 µM biotin-VAD-fmk for an additional 15 min at 37°C. Subsequently, cells were lysed in buffer containing 20 µM biotin-VAD (lanes 1-5). A portion of lysates were then incubated with avidin-Sepharose beads to precipitate active caspases. Immunoblot analysis was performed on precipitates for caspase-8 (top panel), caspase-3 (middle panel), or RIP1 (bottom panel) to reveal active caspases and RIP1 cleavage products. As a positive control for the presence and size of caspase-8, caspase-3, and RIP1, a portion of nonprecipitated whole cell lysates (WCLs) was included in the analysis from fresh (lane 1) or day 4 effectors (lane 2). B, Active caspases were precipitated from day 4 effector lysates and analyzed by immunoblot for the indicated molecules. WCLs were included as a reference comparison. A longer exposure of the caspase-8 immunoblot is included to illustrate precipitation of p43/41 caspase-8. C, Day 4 effectors were treated with FasL for 1 or 2 h, followed by biotin-VAD for 15 min. Active caspases were then precipitated and immunoblot analysis was performed for caspase-8. A portion of the fraction not binding to avidin-Sepharose (Flow Through) as well as WCLs were included in analysis. Bands were revealed using anti-caspase-8 Ab (top panel) or HRP-conjugated Strepavidin (middle and bottom panels). Bands at 55 and 43/41 kDa were identified, which corresponds with the bands revealed in the caspase-8 immunoblots (top and middle panels). In addition, a band of 17 kDa was observed that corresponds with cleaved p17 caspase-3 (bottom panel).

 
A conformational change in procaspase-8 induced by c-FLIPL interaction results in an active full-length protease (10, 18, 27). Distinguishing active full-length caspase-8 from inactive procaspase-8 is thus not possible by standard immunoblot analysis of whole cell lysates. To better identify the degree of caspase activation and the size of active caspases, we used biotin-VAD-fmk to selectively bind enzymatically active caspase catalytic pockets. Fresh day 0 or day 4 effector CD4+ T cells were incubated with biotin-VAD. Lysates were then incubated with avidin-Sepharose beads to affinity purify active caspases. Active caspase-8 and caspase-3 levels were identified using specific Abs in immunoblot analysis of precipitates (Fig. 3A). Whole cell lysates were included to positively identify each caspase band and assess their overall expression (Fig. 3A, lanes 1 and 2). This analysis revealed that full-length caspase-8 bound biotin-VAD in effector T cells (Fig. 3A, top panel, lane 4), but not in unstimulated cells (Fig. 3A, lane 3). A longer exposure of the caspase-8 immunoblot revealed additional p43/41 caspase-8 bands, indicating that this species is also enzymatically active (Fig. 3B, right panel). The binding of biotin-VAD to caspases in effector cell lysates was effectively competed by the addition of 10-fold excess nonbiotinylated z-VAD before incubation of cultures with biotin-VAD (Fig. 3A, lane 5). In addition, no detectable caspase-8 was precipitated using only nonbiotinylated z-VAD (data not shown).

As with caspase-8, more active caspase-3 was present in effector cell lysates than in fresh lysates (Fig. 3A, middle panel, lanes 3 and 4). In contrast with caspase-8, however, there was negligible binding of biotin-VAD to full-length caspase-3, but considerable binding to the small amount of the cleaved p12/10 fragments present in day 4 effector T cell lysates (Fig. 3A, middle panel, lane 4). As with caspase-8, competition with nonbiotinylated z-VAD decreased the amount of precipitated active caspase-3 (Fig. 3A, middle panel, lane 5). These findings are consistent with the accepted model that upstream caspases (e.g., caspase-8) can be active in their full-length form, whereas downstream effector caspases (e.g., caspase-3) require cleavage and dimerization to be active (9, 32). The results demonstrate that activated nonapoptotic CD4+ T cells possess more caspase activity than do unstimulated CD4+ T cells.

Of additional interest was that death domain kinase RIP1 (Fig. 3A, bottom panel, lane 4), but not Fas-associated death domain protein (data not shown), was found to be associated with active caspases precipitated from effector cells. RIP1 is involved with the generation of apoptotic signals by TNFR2 (33) and Fas (34, 35), as well as with the activation of NF-{kappa}B in response to TNF-{alpha} (36, 37), and thus acts as a link between NF-{kappa}B activation and apoptosis. RIP1 both associates with c-FLIP and is a known caspase-8 substrate (33, 38, 39). Reinforcing the observation that RIP1 was associating with active caspases, RIP1 was not seen in precipitates from fresh T cells (Fig. 3A, bottom panel, lane 1). RIP1 association was specifically reduced upon the addition of excess nonbiotinylated z-VAD (Fig. 3A, bottom panel, lane 5). In addition to RIP1, both full-length c-FLIPL and p43FLIP were also precipitated with the active caspase complex (Fig. 3B). Thus, RIP1 and c-FLIP become complexed with active caspases.

To illustrate that p55 caspase-8 is in fact active, effector cells were treated with FasL for 1 or 2 h, and active caspases were labeled, precipitated, and analyzed by immunoblot (Fig. 3C). Whole cell lysates and the portion of cell lysates that did not bind to avidin-Sepharose ("Flow Through") were also included in the immunoblot. As expected, anti-caspase-8 Ab revealed that p55 and p43 caspase-8 were precipitated with biotin-VAD (Fig. 3C, top panel). It is important to note that the proportion of total caspase-8 that was precipitated as active caspase-8 was relatively low, as most caspase-8 in effector T cells did not bind avidin-Sepharose (Fig. 3C, Flow Through). To confirm that biotin-VAD directly bound a 55-kDa species rather than merely coimmunoprecipitating p55 caspase-8, we visualized protein species that bound biotin-VAD using Strepavidin-HRP (Fig. 3C, middle panel) and anti-biotin (data not shown). This also identified bands at 55 kDa and 43/41 kDa, corresponding with full-length caspase-8 and p43/41 caspase-8. In addition, the vast majority of biotin-labeled caspase-8 was precipitated by avidin-Sepharose, as only a very weak band can be visualized in the flow-through fraction that did not bind avidin-Sepharose. An additional 17-kDa band was visualized in whole cell lysates, was enriched with Strepavidin-Sepharose beads, and was absent in the flow-through fractions (Fig. 3C, bottom panel). This band likely corresponds with p17 caspase-3. Thus, biotin-VAD bound directly to full-length p55 caspase-8 in effector but not fresh naive T cells, and this represented only a small fraction of the total caspase-8 in the effector cells.

Caspase blockade inhibits CD4+ T cell proliferation

Given the caspase activity of cycling effector CD4+ T cells, we investigated whether this activity was necessary to initiate cell cycling. CD4+ T cells were labeled with the membrane dye CFSE and stimulated in vitro with anti-CD3/CD28 plus IL-2 in the presence of z-VAD or DMSO vehicle control added at the time of activation. As shown in Fig. 4, both medium control and DMSO-treated cells cycled to a similar degree (36%) by day 2, whereas only 20% of z-VAD-treated cells underwent cell cycling. By day 3, 60% of medium control cells cycled and 58% of the DMSO control cells, compared with only 32% of cells that received z-VAD. These results indicate that a blockade in caspase activation prevents cycling of murine T cells.



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  		FIGURE 4. z-VAD inhibits CD4+ T cell proliferation. z-VAD (100 µM) or a comparable volume of DMSO vehicle control was added to CFSE-labeled polyclonal CD4+ T cell cultures at day 0 and stimulated with anti-CD3/CD28 in the presence of IL-2. CD4+ T cells were analyzed for proliferation as measured by CFSE dye loss on day 2 (top panel) or 3 (bottom panel). Number insets represent the percentage of cells that divided at least one time.

 
Th2 effectors manifest more caspase activity than do Th1 effectors

Given the role of caspase activity for both cell growth and cell death, the level of caspase activity in effector T cells could profoundly influence the proportion of various effector subsets. As Th1 and Th2 cells are differentially responsive to Fas-mediated cell death, we characterized caspase activity and cleavage of caspase substrates in these cells (40, 41). We also wished to determine whether our findings using anti-CD3/CD28 stimulation of polyclonal CD4+ T cells extended to Ag-specific CD4+ T cells. For this purpose we used CD4+ T cells from AND TCR transgenic mice, which recognize PCCF presented in the context of I-EK (42).

AND Th1 and Th2 effector CD4+ T cells were produced in vitro, as described in Materials and Methods. Intracellular cytokine staining and ELISA were used to verify correct polarization of the resulting day 4 effectors (data not shown). Viable day 4 effector T cells were purified by Percoll centrifugation, and a portion was then used in the DEVD-rhodamine release assay to determine overall caspase activity (Fig. 5A). AND Th2 effectors had significantly higher overall caspase activity than did Th1 effectors. The remaining day 4 effectors were extensively washed and rested for 3 days in the absence of exogenous cytokines to reduce cell cycling (43), and viable rested effectors were then purified. Concomitant with their reduced cell cycling, rested effectors manifested considerably diminished caspase activity (Fig. 5B). Nonetheless, Th2 rested effector cells possessed higher caspase activity than did Th1 rested effector cells (Fig. 5B, inset). Thus, caspase activity closely paralleled cell cycling.



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  		FIGURE 5. AND Th2 CD4+ T cells possess higher levels of caspase activity than do Th1 cells at the effector and rested effector stages. AND naive CD4+ T cells were stimulated in vitro under polarizing conditions for 4 days using DCEK-ICAM APCs pulsed with 5 µM PCCF. Viable cells were used to perform caspase activity assays. Shown is a representative example of three experiments. A, Caspase activity of day 4 effectors. Results are representative of three experiments (p = 0.0044, two-way ANOVA column factor). B, Portions of Th1 and Th2 effector populations were washed three times to remove exogenous cytokines and were cultured in the absence of exogenous cytokines for an additional 3 days to generate rested effector populations (43 ). Results are displayed using the same scale as in A. Figure inset in B represents results from the same rested effectors using a more sensitive scale that shows residual persistent caspase activity to be more prominent in Th2 rested effectors.

 
To better define which caspases were active, and to what degree, in Th1 and Th2 effectors, biotin-VAD was again used to purify active caspases. In agreement with the DEVD-rhodamine caspase activity assay, Th2 effectors possessed more active full-length caspase-8 than did Th1 effectors (Fig. 6, top panel, lanes 3 and 4), despite having identical overall levels and cleavage patterns of caspase-8 in their whole cell lysates. Comparable results were seen with analysis of active caspase-3 (Fig. 6, middle panel). In addition, more RIP1 was seen in complex with active caspases in Th2 cells than in Th1 cells (Fig. 6, bottom panel).



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  		FIGURE 6. Th2 effector CD4+ T cells possess more active full-length caspase-8, active cleaved caspase-3, and RIP1 in complex with active caspases than do Th1 effectors. Day 4 Th1 and Th2 effector CD4+ T cells were generated using anti-CD3/CD28 stimulation and then were incubated at 37°C for 15 min with biotin-VAD followed by lysis in buffer containing 20 µM biotin-VAD. A portion of each lysate was incubated with avidin-Sepharose beads to isolate active caspases. Immunoblot analysis was performed on nonprecipitated WCLs (lanes 1 and 2) and precipitates (lanes 3 and 4) for caspase-8 (top panel), caspase-3 (middle panel), or RIP1 (bottom panel). NS, Nonspecific band.

 
To determine the downstream targets of increased caspase activity in Th2 effectors, we examined cleavage patterns of candidate caspase substrates in viable AND naive cells as well as in viable polarized AND CD4+ effector and rested effector T cells stimulated with PCCF. AND naive CD4+ T cells contained predominantly full-length c-FLIPL, whereas activated day 4 Th1 and Th2 effectors expressed cleaved p43FLIP in addition to full-length c-FLIPL (Fig. 7A). No difference was observed in FLIP RNA levels between Th1 and Th2 effectors (data not shown). However, Th2 cells possessed an additional p27FLIP product with proportionally less full-length c-FLIPL and p43FLIP. The p27FLIP fragment that we observed was of a molecular mass similar to that of an alternative splice product of FLIP message (FLIPshort), which has been reported to be up-regulated in activated T cells and during restimulation events (44, 45). The p27FLIP fragment that we observed corresponds with a second putative caspase cleavage site at Asp258. In this instance, it would appear that p27FLIP was more likely a further cleavage product than an alternative splice form. Evidence for this was observed in CD4+ T cell lysates from full-length c-FLIPL transgenic mice. In the case of the transgenic animal, the transgene cannot be alternatively spliced (Fig. 7A). This finding is also consistent with the observation of elevated caspase activity in T cells from c-FLIPL transgenic mice (R. C. Budd, unpublished observations). Consistent with the concept that p27FLIP can be generated by active caspases, wild-type CD4+ T cells treated with FasL also manifest a p27FLIP fragment (Fig. 2B). Rested effectors showed a return of full-length c-FLIPL (Fig. 7A), consistent with the observed decrease in the caspase activity of these cells (Fig. 5). Nonetheless, the residual higher level of caspase activity in Th2 rested effectors (Fig. 5) was reflected in an increased amount of p27FLIP compared with that seen in Th1 rested effector cells.



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  		FIGURE 7. AND Th2 effector CD4+ T cells show more extensive cleavage of selective upstream caspase substrates than do Th1 effectors. Viable naive (Fresh) day 4 Ag-activated Th1 and Th2 effectors and Th1 and Th2 rested effector populations were purified, and immunoblot analysis of lysates was performed for c-FLIPL (A), RIP1 (B), Bid (C), and ICAD (D). Lysates from c-FLIPL transgenic CD4+ T cells were included in A to demonstrate the p43FLIP and p27FLIP cleavage fragments of overexpressed full-length c-FLIPL. B, RIP1 represents noncleaved protein and p38RIP1 represents a cleavage product generated by cleavage at Asp324. NS, Nonspecific band. C, t-BID is generated by caspase-mediated cleavage at Asp75. D, Cleavage of ICAD at Asp224 results in p18ICAD. Lysates of RIP1–/– 3T3 fibroblasts were included as negative controls. Bid (C) and ICAD (D) remained as mostly noncleaved proteins under these culture conditions. Results are representative of four experiments.

 
To assess the extent of propagation of the caspase cascade in polarized effector T cells, we examined cleavage of the caspase-8 substrates RIP1 and Bid and the downstream caspase-3 substrate ICAD (Fig. 7, B–D). Given our findings with c-FLIP cleavage, RIP1 was a particularly attractive candidate as it both associates with c-FLIP and is a known caspase-8 substrate, yielding a p38RIP1 fragment (33, 38). We observed that RIP1 cleavage patterns in Th1 and Th2 effectors mimicked that of c-FLIPL. AND naive CD4+ T cells expressed mainly full-length RIP1, whereas effector cells manifested the p38RIP1 cleavage product with concomitant reduction of full-length RIP1 (Fig. 7B). In each sample, full-length RIP1 appeared as three distinct bands, consistent with recent reports that RIP1 can be ubiquitylated (46). Th2 effectors possessed less full-length RIP1 than did Th1 effectors (Fig. 7B). As with c-FLIP cleavage products, levels of p38RIP1 diminished in both Th1 and Th2 rested effector cells, but Th2 rested effectors manifested less full-length RIP1 (Fig. 7B). The specificity of the full-length RIP1 and p38RIP1 bands was confirmed using lysates from both RIP1-deficient 3T3 cells as a negative control (Fig. 7B).

Unlike c-FLIPL and RIP1, Bid is not known to form a complex with caspase-8, although Bid can be cleaved by caspase-8. Interestingly, in contrast with c-FLIPL and RIP1, Bid showed only minimal levels of cleavage in either polarized effector or rested effector populations (Fig. 7C). This finding closely paralleled the earlier observations with nonpolarized CD3/CD28-stimulated effector cells (Fig. 2C). In addition, no observed cleavage of the caspase-3 substrate ICAD was detected in either Th1 or Th2 effector or rested effector populations. Collectively, these data show that Th2 cells manifest greater caspase activity and cleavage of upstream caspase substrates than do Th1 cells at the effector and rested effector stages.

Th2 effectors may better tolerate elevated caspase activity due to higher levels of Bcl-2

Conceivably, the higher levels of caspase activity of Th2 cells might result in greater cell death over Th1 cells. However, TUNEL analysis of polarized Th1 and Th2 effector cells actually revealed higher levels of apoptosis in Th1 effectors on days 3 and 4 (Fig. 8A). Similar results were found in polarized polyclonal CD4+ T cell populations (data not shown). The differences between TUNEL+ cells in Th1 and Th2 populations on day 4, but not day 3, reached statistical significance (p < 0.05, n = 2). This finding is consistent with an earlier report that a higher proportion of Th1 cells die compared with Th2 effectors (40).



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  		FIGURE 8. AND Th1 effector CD4+ T cells exhibit a greater population of Bcl-2– and dead cells than do Th2 cells. A, TUNEL analysis of day 3 and 4 AND effector CD4+ T cells stimulated with APCs and PCCF. Data represent mean ± SD from triplicate cultures. B, Cell death (nicked DNA) was assessed by TUNEL assay and intracellular Bcl-2 levels were determined on day 4 Th1 and Th2 effectors in which 100 µM z-VAD, or equivalent volume of DMSO, was added at day 0. Percentage insets represent the proportion of total cells. Numbers in parentheses represent the proportion of Bcl-2– cells containing nicked DNA. Data are representative of five independent experiments.

 
Because the increased caspase activity in Th2 effectors did not result in increased apoptosis, we considered that there might be a mechanism by which Th2 effectors better tolerate elevated caspase activity. To explore this possibility, we examined expression of a major protector against caspase-driven death, Bcl-2, which functions to protect the outer mitochondrial membrane from caspase-initiated cell death (47). Day 4 Th1 and Th2 effectors were analyzed by simultaneous staining for apoptotic cells by TUNEL assay and for intracellular Bcl-2. Th1 effectors expressed a higher proportion of Bcl-2– cells (23%) than did Th2 effectors (12%) (Fig. 8B). Th2 AND effectors manifest a statistically higher percentage of cells of the phenotype Bcl-2high/TUNEL– and lower levels of Bcl-2low/TUNEL– cells (p < 0.05, n = 2). In addition, the apoptotic cells were almost entirely confined to the Bcl-2– subpopulation, suggesting that the presence of Bcl-2 was protective against the caspase activity in CD4+ effector T cells. This was reinforced by the addition of z-VAD to cultures at day 0, which reduced the proportion of TUNEL-positive cells in the Bcl-2– population from 22% to 15% in Th1 cultures and from 17% to 8% in Th2 cultures (Fig. 8B). Caspase blockade also yielded an overall increase in the proportion of Bcl-2– cells. Th2 effector cells are thus potentially better adapted to sustain higher levels of caspase activity, in part through maintenance of higher levels of Bcl-2 expression.

An additional, though not mutually exclusive, possibility is that Th2 effector cells may have a greater requirement for caspase activity for cell cycling. To examine this, CFSE-labeled AND CD4+ T cells were stimulated with PCCF under Th1 and Th2 polarizing conditions with or without z-VAD added at day 0 of culture. Equivalent fractions of Th1 (10%) and Th2 (7%) cells proliferated by day 2 in the absence of caspase blockade (Fig. 9, DMSO). However, caspase blockade reduced the percentage of proliferating cells to 1% in both the Th1 and Th2 populations. By day 4, 94% of Th1 and 95% of Th2 cells had proliferated, whereas caspase blockade at day 0 decreased proliferation to 58% in Th1 and 55% in Th2 cells. Furthermore, a delay in the addition of z-VAD until day 2 had no effect on subsequent proliferation by AND CD4+ T cells (data not shown). These findings suggest that caspase activation is equally necessary for initial activation of the majority of Th1 and Th2 populations, but that it is not required for continued proliferation during the Ag-independent, cytokine-driven phase of CD4+ T cell growth (20).



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  		FIGURE 9. Addition of z-VAD inhibits T cell proliferation under either Th1 or Th2 polarization conditions. CFSE-labeled AND CD4+ T cells were cultured under Th1 or Th2 polarizing conditions in the presence of 100 µM z-VAD or appropriate DMSO control added at day 0 and then were analyzed for cell cycling on days 2 (top panel) and 4 (bottom panel). Number insets represent the percentage of divided cells.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Our results show that viable effector CD4+ T cells generate caspase activity levels that are between those of fresh CD4+ T cells and FasL-treated effector T cells. This is reflected in the cleavage of caspase-8 substrates c-FLIPL, RIP1, and to a lesser extent Bid, but not the downstream caspase-3 substrate ICAD. In addition, RIP1, full-length c-FLIPL, and p43FLIP were precipitated with active caspases in effector T cells. Furthermore, this intermediate caspase activity is necessary for the initiation of cell cycling by the majority of T cells. Caspase activity is also higher in Th2 vs Th1 viable effector CD4+ T cells and subsides in each population as they decrease cell cycling and become rested effector cells. Activated T cells thus run a delicate balance of generating sufficient caspase activity to promote cell cycling, but not so much as to induce cell death. The levels of caspase activity may profoundly affect the population of T cell effector subsets.

Because there is a fine balance of caspase activity between proliferation and cell death, it is important to restrain and target caspase activity to specific substrates in living cells. The recent concept that noncleaved caspase-8 can manifest intermediate activity is fully consistent with our results (18). Our findings further suggest that this form of active caspase-8 leads to the cleavage of molecules known to associate with it, including c-FLIPL and RIP1 (39), with less extensive cleavage of more distal substrates such as Bid. This would suggest that molecules that complex with caspase-8 are most efficiently cleaved in effector cells. Although we speculate that RIP1 is binding to caspase-8, it remains formally possible that RIP1 is binding to and cleaved by active caspases other than caspase-8. Furthermore, the limited degree of caspase-3 activity in effector T cells does not lead to any appreciable ICAD cleavage.

Recent reports show that caspase-3 and caspase-8 can localize in lipid rafts of nonactivated peripheral blood lymphocytes and that caspase-3 levels increase in lipid rafts upon activation (48). In addition, these and other investigators have shown that p20 caspase-3 is kept inactive by X-linked inhibitor of apoptosis protein (XIAP) and is activated upon further cleavage in which the prodomain of capase-3 is released, thereby generating p17 caspase-3 (29, 48). If second mitochondria-derived activator of caspase/direct inhibitor of apoptosis-binding protein with low isoelectric point protein is released from the mitochondria, then XIAP is inactivated, thereby enabling the autoproteolytic processing of p20 caspase-3 to p17 caspase-3. Bcl-2 protein present in the mitochondria inhibits release of second mitochondria-derived activator of caspase, thereby maintaining the association of XIAP and p20 caspase-3 and preventing further processing of caspase-3 (29, 30, 31). Studies have suggested that other active caspases, such as caspase-8 or caspase-9, can process caspase-3 to its fully mature 17-kDa fragment (49). Thus, the likely need to maintain mitochondrial integrity to prevent excessive caspase activity at the effector T cell stage is reflected in the high percentage of apoptotic cells in the Bcl-2– subset. In agreement with this concept, caspase blockade allowed greater survival of Bcl-2– cells.

Further support for the notion that p20 caspase-3 is inactive derives from its presence in freshly isolated T cells, in which negligible levels of caspase activity were detected. However, in day 4 effector T cells, where caspase activity was detected, the intensity of the p20 caspase-3 band decreased concomitant with an increase of detectable p12/10 caspase-3. By day 6, there was a decrease in p12/10 caspase-3 with a proportional increase in p20 caspase-3. Taken together, these results suggest that p20 caspase-3 is likely inactive, consistent with it being complexed with XIAP (29). Furthermore, our results showing negligible binding of biotin-VAD to either full-length pro-caspase-3 or p20 caspase-3, but binding to a 17-kDa fragment, are consistent with this view and with other studies (29, 48).

It was recently reported that c-FLIPS, a 27-kDa alternate splice variant of c-FLIP, blocks caspase-8 activation to a greater extent than does FLIPL (28). Both c-FLIPL and c-FLIPS have been reported to inhibit TRAIL receptor-mediated apoptosis (50, 51, 52, 53, 54). c-FLIPS protein is up-regulated within the first day of activation of primary human T cells (45). When cyclohexamide is used during this time frame, FLIPS is down-regulated and cells are sensitive to Fas-mediated mitochondrial membrane potential loss, despite normal expression of Bcl-2 family members (45). Levels of c-FLIPS diminish over time and by day 6 are nearly undetectable (45). Other studies show that restimulation of day 6 effectors with anti-CD3/CD-28 also leads to up-regulation of c-FLIPS protein and partial protection of mitochondrial integrity, compared with restimulation with only anti-CD3 (44). In both studies, increased c-FLIPS protein expression correlated with decreased caspase-8 processing at the Fas-mediated death-inducing signaling complex (44, 45). Other studies indicate a role for elevated c-FLIPS in the survival of various tumor cells, including human gastric cancers (55, 56) and cells from patients with myelodysplastic syndrome (57). We propose an alternative mechanism of generating p27FLIP by further cleavage of p43FLIP. In addition to the known caspase cleavage site at Asp376 (LEVD), there is a second potential caspase cleavage site at Asp258 (LIID) that would lead to p27FLIP. We found that p27FLIP is also a cleavage product in T cell lysates from c-FLIPL transgenic mice as well as from FasL treatment of T cell blasts. The fact that Th2 cells possess more caspase activity and greater p27FLIP than do Th1 cells is also consistent with proteolytic generation of this fragment. In addition, quantitative PCR revealed that c-FLIP message is expressed at very similar levels in Th1 and Th2 cells (data not shown). If the p27FLIP that we observed were to be generated via an alternative splicing mechanism, then we would expect to see an increase in c-FLIP message in Th2 cells. p27FLIP may thus arise by two mechanisms in activated T cells.

Our results expand on other recent studies showing a requirement for caspases in T cell function. In addition, human T cells bearing a mutation in the caspase-8 gene exhibited decreased IL-2 production and CD25 induction upon stimulation (7). Restoration of wild-type caspase-8 in these cells promoted normal induction of CD25 upon TCR stimulation (7). Consistent with these findings, inhibition of caspase-8 expression by siRNA decreased activation of T cells (7). In contrast, caspase-8 conditional knockout mice did not show a defect in IL-2 production, but did exhibit a defect in response to IL-2, which could not be rescued by PMA and ionomycin (8). Thus, multiple studies highlight the importance of caspases in T cell effector function. Yet, it has been uncertain just how much caspase activity is present in cycling vs apoptotic T cells, what the substrates are, and how extensive is the progression of the caspase cascade.

It is not presently clear why Th2 cells exhibit higher caspase activity than do Th1 cells. One potential explanation is that Th2 effectors are better equipped to survive caspase activation. This possibility is supported by the findings of a higher percentage of Bcl-2+ cells among Th2 cells compared with Th1 effectors. These results may offer a partial explanation for two reports that find that Th1 cells are more susceptible to Fas-mediated cell death (40, 41). Alternatively, though not mutually exclusively, Th2 cells may require higher levels of caspase activity to obtain their fully polarized cytokine phenotype by cleavage of selective substrates important in signal transduction pathways. Studies to explore the latter possibility are currently underway.

The exact contribution of caspase activity and their substrates in T cell activation remains undefined, though the current findings identify logical candidate substrates. One role for caspases could be to cleave c-FLIPL to its p43 form, which more efficiently recruits RIP1 and TNFR-associated factor 2, leading to activation of NF-{kappa}B (58). Consistent with this hypothesis, we observed that RIP1 was in a complex with and proportional to levels of active caspases and overall p43FLIP levels (Figs. 3 and 6). Several studies have revealed a role of Fas signaling in neurite outgrowth (1), fibroblast proliferation (4), maturation of dendritic cells (5), and hepatocyte regeneration (59). Conceivably, Fas may engage signal pathways involved with cell growth, such as Erk activation (1). However, we currently have no evidence that Fas is required for the initiation of caspase activity in effector T cells. In fact, preliminary studies show similar caspase activity is generated in Fas-deficient lpr effector T cells (J. Q. Russell, unpublished observations).

The direct connection between TCR ligation and activation of caspases has not yet been identified. A logical consideration is that expression of a death receptor ligand is up-regulated, which stimulates its cognate receptor to activate caspases. This is consistent with previous findings showing that Fas ligation can costimulate T cell proliferation (6, 21, 60). However, as mentioned above, Fas-deficient lpr T cells proliferate normally and activate caspases upon TCR stimulation. An alternative view is that TCR activation may promote approximation of caspase-8 with c-FLIPL independently of a death receptor, possibly via lipid raft association as suggested for Fas signaling in activated T cells (61). In this regard, c-FLIPL transgenic mouse CD4+ T cells manifest increased caspase activity (R. C. Budd, unpublished observations) and hyperproliferate in response to TCR engagement (62).

Caspases (especially caspase-8) and c-FLIPL clearly have an important role in the growth and development of potentially many cell types. This is underscored by the cardiac developmental block in mice deficient for either c-FLIPL (63) or caspase-8 (64). Defining the substrates of caspases that are critical for cell growth will represent an important link between the mechanisms of cell growth and cell death and may suggest an important central process for preventing aberrant growth by linking cell cycling to the same pathways used at a higher intensity for cell death.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Acknowledgments
 
We thank Colette Charland for technical assistance with flow cytometry, members of the University of Vermont DNA facility for assistance with real-time PCR, and University of Vermont Immunobiology Program members for helpful discussions and suggestions.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by grants from the National Institutes of Health (AI36333 and AI45666 to R.C.B.; AI26887 and AI46530 to S.L.S.). R.S.M. is supported by National Institutes of Health Grant No. 5T32ES007122. Back

2 Address correspondence and reprint requests to Dr. Ralph C. Budd, University of Vermont, Immunobiology Program, College of Medicine, 89 Beaumont Avenue, Given Building D305, Burlington, VT 05405. E-mail address: ralph.budd{at}uvm.edu Back

3 Abbreviations used in this paper: ICAD, inhibitor of caspase-activated DNase; RIP1, receptor interacting protein 1; PCCF, pigeon cytochrome c fragment; z-VAD, z-Val-Ala-Asp-fluoromethylketone; FasL, Fas ligand; WCL, whole cell lysate; XIAP, X-linked inhibitor of apoptosis protein. Back

Received for publication July 14, 2004. Accepted for publication January 11, 2005.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Desbarats, J., R. B. Birge, M. Mimouni-Rongy, D. E. Weinstein, J. S. Palerme, M. K. Newell. 2003. Fas engagement induces neurite growth through ERK activation and p35 upregulation. Nat. Cell Biol. 5:118.[Medline]
  2. Wollert, K. C., J. Heineke, J. Westermann, M. Ludde, B. Fiedler, W. Zierhut, D. Laurent, M. K. Bauer, K. Schulze-Osthoff, H. Drexler. 2000. The cardiac Fas (APO-1/CD95) Receptor/Fas ligand system: relation to diastolic wall stress in volume-overload hypertrophy in vivo and activation of the transcription factor AP-1 in cardiac myocytes. Circulation 101:1172.[Abstract/Free Full Text]
  3. Desbarats, J., M. K. Newell. 2000. Fas engagement accelerates liver regeneration after partial hepatectomy. Nat. Med. 6:920.[Medline]
  4. Aggarwal, B. B., S. Singh, R. LaPushin, K. Totpal. 1995. Fas antigen signals proliferation of normal human diploid fibroblast and its mechanism is different from tumor necrosis factor receptor. FEBS Lett. 364:5.[Medline]
  5. Rescigno, M., V. Piguet, B. Valzasina, S. Lens, R. Zubler, L. French, V. Kindler, J. Tschopp, P. Ricciardi-Castagnoli. 2000. Fas engagement induces the maturation of dendritic cells (DCs), the release of interleukin (IL)-1{beta}, and the production of interferon {gamma} in the absence of IL-12 during DC-T cell cognate interaction: a new role for Fas ligand in inflammatory responses. J. Exp. Med. 192:1661.[Abstract/Free Full Text]
  6. Alderson, M. R., R. J. Armitage, E. Maraskovsky, T. W. Tough, E. Roux, K. Schooley, F. Ramsdell, D. H. Lynch. 1993. Fas transduces activation signals in normal human T lymphocytes. J. Exp. Med. 178:2231.[Abstract/Free Full Text]
  7. Chun, H. J., L. Zheng, M. Ahmad, J. Wang, C. K. Speirs, R. M. Siegel, J. K. Dale, J. Puck, J. Davis, C. G. Hall, et al 2002. Pleiotropic defects in lymphocyte activation caused by caspase-8 mutations lead to human immunodeficiency. Nature 419:395.[Medline]
  8. Salmena, L., B. Lemmers, A. Hakem, E. Matysiak-Zablocki, K. Murakami, P. Y. Au, D. M. Berry, L. Tamblyn, A. Shehabeldin, E. Migon, et al 2003. Essential role for caspase 8 in T-cell homeostasis and T-cell-mediated immunity. Genes Dev. 17:883.[Abstract/Free Full Text]
  9. Degterev, A., M. Boyce, J. Yuan. 2003. A decade of caspases. Oncogene 22:8543.[Medline]
  10. Muzio, M., B. R. Stockwell, H. R. Stennicke, G. S. Salvesen, V. M. Dixit. 1998. An induced proximity model for caspase-8 activation. J. Biol. Chem. 273:2926.[Abstract/Free Full Text]
  11. Lavrik, I., A. Krueger, I. Schmitz, S. Baumann, H. Weyd, P. H. Krammer, S. Kirchhoff. 2003. The active caspase-8 heterotetramer is formed at the CD95 DISC. Cell Death Differ. 10:144.[Medline]
  12. Enari, M., H. Sakahira, H. Yokoyama, K. Okawa, A. Iwamatsu, S. Nagata. 1998. A caspase-activated DNase that degrades DNA during apoptosis, and its inhibitor ICAD. Nature 391:43.[Medline]
  13. Mashima, T., M. Naito, K. Noguchi, D. K. Miller, D. W. Nicholson, T. Tsuruo. 1997. Actin cleavage by CPP-32/apopain during the development of apoptosis. Oncogene 14:1007.[Medline]
  14. Martin, S. J., D. M. Finucane, G. P. Amarante-Mendes, G. A. O’Brien, D. R. Green. 1996. Phosphatidylserine externalization during CD95-induced apoptosis of cells and cytoplasts requires ICE/CED-3 protease activity. J. Biol. Chem. 271:28753.[Abstract/Free Full Text]
  15. Lazebnik, Y. A., A. Takahashi, G. G. Poirier, S. H. Kaufmann, W. C. Earnshaw. 1995. Characterization of the execution phase of apoptosis in vitro using extracts from condemned-phase cells. J. Cell Sci. Suppl. 19:41.[Medline]
  16. Liu, X., C. N. Kim, J. Yang, R. Jemmerson, X. Wang. 1996. Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell 86:147.[Medline]
  17. Rodriguez, J., Y. Lazebnik. 1999. Caspase-9 and APAF-1 form an active holoenzyme. Genes Dev. 13:3179.[Abstract/Free Full Text]
  18. Micheau, O., M. Thome, P. Schneider, N. Holler, J. Tschopp, D. W. Nicholson, C. Briand, M. G. Grutter. 2002. The long form of FLIP is an activator of caspase-8 at the Fas death-inducing signaling complex. J. Biol. Chem. 277:45162.[Abstract/Free Full Text]
  19. Scaffidi, C., I. Schmitz, P. H. Krammer, M. E. Peter. 1999. The role of c-FLIP in modulation of CD95-induced apoptosis. J. Biol. Chem. 274:1541.[Abstract/Free Full Text]
  20. Jelley-Gibbs, D. M., N. M. Lepak, M. Yen, S. L. Swain. 2000. Two distinct stages in the transition from naive CD4 T cells to effectors, early antigen-dependent and late cytokine-driven expansion and differentiation. J. Immunol. 165:5017.[Abstract/Free Full Text]
  21. Kennedy, N. J., T. Kataoka, J. Tschopp, R. C. Budd. 1999. Caspase activation is required for T cell proliferation. J. Exp. Med. 190:1891.[Abstract/Free Full Text]
  22. Alam, A., L. Y. Cohen, S. Aouad, R. P. Sekaly. 1999. Early activation of caspases during T lymphocyte stimulation results in selective substrate cleavage in nonapoptotic cells. J. Exp. Med. 190:1879.[Abstract/Free Full Text]
  23. Garcia-Calvo, M., E. P. Peterson, B. Leiting, R. Ruel, D. W. Nicholson, N. A. Thornberry. 1998. Inhibition of human caspases by peptide-based and macromolecular inhibitors. J. Biol. Chem. 273:32608.[Abstract/Free Full Text]
  24. Wei, Y., T. Fox, S. P. Chambers, J. Sintchak, J. T. Coll, J. M. Golec, L. Swenson, K. P. Wilson, P. S. Charifson. 2000. The structures of caspases-1, -3, -7 and -8 reveal the basis for substrate and inhibitor selectivity. Chem. Biol. 7:423.[Medline]
  25. Shi, Y.. 2002. Mechanisms of caspase activation and inhibition during apoptosis. Mol. Cell 9:459.[Medline]
  26. Irmler, M., M. Thome, M. Hahne, P. Schneider, K. Hofmann, V. Steiner, J.-L. Bodmer, M. Schroter, K. Burns, C. Mattmann, et al 1997. Inhibition of death receptor signals by cellular FLIP. Nature 388:190.[Medline]
  27. Chang, D. W., Z. Xing, Y. Pan, A. Algeciras-Schimnich, B. C. Barnhart, S. Yaish-Ohad, M. E. Peter, X. Yang. 2002. c-FLIPL is a dual function regulator for caspase-8 activation and CD95-mediated apoptosis. EMBO J. 21:3704.[Medline]
  28. Krueger, A., I. Schmitz, S. Baumann, P. H. Krammer, S. Kirchhoff. 2001. Cellular FLICE-inhibitory protein splice variants inhibit different steps of caspase-8 activation at the CD95 death-inducing signaling complex. J. Biol. Chem. 276:20633.[Abstract/Free Full Text]
  29. Sun, X. M., S. B. Bratton, M. Butterworth, M. MacFarlane, G. M. Cohen. 2002. Bcl-2 and Bcl-xL inhibit CD95-mediated apoptosis by preventing mitochondrial release of Smac/DIABLO and subsequent inactivation of X-linked inhibitor-of-apoptosis protein. J. Biol. Chem. 277:11345.[Abstract/Free Full Text]
  30. Fernandes-Alnemri, T., R. C. Armstrong, J. Krebs, S. M. Srinivasula, L. Wang, F. Bullrich, L. C. Fritz, J. A. Trapani, K. J. Tomaselli, G. Litwack, E. S. Alnemri. 1996. In vitro activation of CPP32 and Mch3 by Mch4, a novel human apoptotic cysteine protease containing two FADD-like domains. Proc. Natl. Acad. Sci. USA 93:7464.[Abstract/Free Full Text]
  31. Stennicke, H. R., J. M. Jurgensmeier, H. Shin, Q. Deveraux, B. B. Wolf, X. Yang, Q. Zhou, H. M. Ellerby, L. M. Ellerby, D. Bredesen, et al 1998. Pro-caspase-3 is a major physiologic target of caspase-8. J. Biol. Chem. 273:27084.[Abstract/Free Full Text]
  32. Salvesen, G. S., V. M. Dixit. 1997. Caspases: intracellular signaling by proteolysis. Cell 91:443.[Medline]
  33. Lin, Y., A. Devin, Y. Rodriguez, Z. G. Liu. 1999. Cleavage of the death domain kinase RIP by caspase-8 prompts TNF-induced apoptosis. Genes Dev. 13:2514.[Abstract/Free Full Text]
  34. Holler, N., R. Zaru, O. Micheau, M. Thome, A. Attinger, S. Valitutti, J. L. Bodmer, P. Schneider, B. Seed, J. Tschopp. 2000. Fas triggers an alternative, caspase-8-independent cell death pathway using the kinase RIP as effector molecule. Nat. Immunol. 1:489.[Medline]
  35. Stanger, B. Z., P. Leder, T. H. Lee, E. Kim, B. Seed. 1995. RIP: a novel protein containing a death domain that interacts with Fas/APO-1 (CD95) in yeast and causes cell death. Cell 81:513.[Medline]
  36. Ting, A. T., F. X. Pimentel-Muinos, B. Seed. 1996. RIP mediates tumor necrosis factor receptor 1 activation of NF-{kappa}B but not Fas/APO-1-initiated apoptosis. EMBO J. 15:6189.[Medline]
  37. Kelliher, M. A., S. Grimm, Y. Ishida, F. Kuo, B. Z. Stanger, P. Leder. 1998. The death domain kinase RIP mediates the TNF-induced NF-{kappa}B signal. Immunity 8:297.[Medline]
  38. Martinon, F., N. Holler, C. Richard, J. Tschopp. 2000. Activation of a pro-apoptotic amplification loop through inhibition of NF-{kappa}B-dependent survival signals by caspase-mediated inactivation of RIP. FEBS Lett. 468:134.[Medline]
  39. Kataoka, T., R. C. Budd, N. Holler, M. Thome, F. Martinon, M. Irmler, K. Burns, M. Hahne, N. Kennedy, M. Kovacsovics, J. Tschopp. 2000. The caspase-8 inhibitor FLIP promotes activation of NF-{kappa}B and Erk signaling pathways. Curr. Biol. 10:640.[Medline]
  40. Zhang, X., T. Brunner, L. Carter, R. W. Dutton, P. Rogers, L. Bradley, T. Sato, J. C. Reed, D. Green, S. L. Swain. 1997. Unequal death in T helper cell (Th)1 and Th2 effectors: Th1, but not Th2, effectors undergo rapid Fas/FasL-mediated apoptosis. J. Exp. Med. 185:1837.