CTL are endowed with the ability to eliminate pathogens through perforin-mediated cytotoxic activity. The mechanism for perforin-mediated Ag-specific killing has been solely attributed to cytotoxic granule exocytosis from activated CD8+ T cells. In this study, we redefine this mechanism, demonstrating that virus-specific CD8+ T cells rapidly up-regulate perforin in response to stimulation temporally with IFN-γ and CD107a expression. Following Ag-specific activation, newly synthesized perforin rapidly appears at the immunological synapse, both in association with and independent of cytotoxic granules, where it functions to promote cytotoxicity. Our work suggests a novel mechanism of CTL cytotoxicity and identifies a novel correlate of CD8+ T cell-mediated immunity.

Cytotoxic T lymphocytes are endowed with the ability to eliminate tumor cells as well as host cells that harbor intracellular pathogens via two principal mechanisms: 1) the exocytosis of preformed secretory granules that contain cytotoxic proteins (1, 2, 3) and 2) the engagement of receptors on the target cell that induce cellular apoptosis, such as those specific for Fas ligand (4, 5), TNF (6), and TRAIL (6). Cytotoxic granules contain various granzymes (7), a family of highly specific serine proteases that initiate apoptosis (8, 9, 10, 11), and perforin (7), a protein capable of binding phospholipid membranes in a calcium-dependent manner (12, 13, 14). During activation, perforin monomers polymerize on lipid bilayers to form a pore (12, 14, 15) that permits entry of granzymes into the target cell to induce apoptosis. How perforin enables granzyme entry is not known, however, as the localization of pore formation within the target cell remains controversial. Nonetheless, in both humans and mice, abrogation of perforin activity results in profoundly impaired cellular cytotoxicity and immunodeficiency (16, 17).

CTL are capable of serial killing, the sequential elimination of several target cells (18, 19), but the mechanism that permits this ability is unclear. CTL may 1) ration their granules, 2) rapidly up-regulate their cytotoxic proteins to sustain/replenish their killing ability, or 3) need to proliferate to recover their cytotoxicity. Cytotoxic granule-mediated killing by CTL occurs within hours of target cell recognition (19), but reconstitution of intracellular perforin has only been detected after several days of proliferation (20, 21, 22). On this premise, the current paradigm is that CTL must proliferate before perforin recovery, implying that the cytotoxic granule content of a resting CTL dictates the immediate killing potential of that cell. Biologically, this seems inefficient, as an infecting virus that could outlast the initial brunt of the effector CD8+ T cell population would continue to replicate in an unfettered manner until the CD8+ T cell population had proliferated. A potential solution to this conundrum is the possibility of rapid recharging of the CTL cytotoxic machinery following activation. Such a mechanism would allow the CTL to continue killing following granule release.

Although rapid perforin up-regulation was shown to occur in NK cell lines (23), this ability in primary CD8+ T cells has until recently been unclear. Contrary to previous reports, we found that perforin up-regulation by Ag-experienced human CD8+ T cells occurs within hours of activation, without a requirement for exogenous cytokines or cellular proliferation (24). However, the role of new perforin remains unknown; is it simply a means to replenish depleted granules or is it also actively used to mediate cytotoxicity? In this study, we provide evidence that newly produced perforin traffics directly to the immunological synapse, where it promotes cytotoxic activity. These results define perforin up-regulation ability as a novel correlate of cytotoxic potential in human CD8+ T cells.

PBMC were obtained from a normal human subject who exhibited an exceptionally strong response to the HLA-B7-restricted human CMV pp65 peptide TM10: TPRVTGGGAM epitope (4235 spot-forming cells per 106 PBMC). Donor PBMC were collected by the University of Pennsylvania’s Center for AIDS Research Human Immunology Core, in compliance with the guidelines set by the institution’s internal review board, and cryopreserved in FBS (HyClone) containing 10% DMSO (Fisher Scientific).

Abs for surface staining included anti-CD4 PE Cy5-5 (Invitrogen), anti-CD107a FITC (BD Biosciences), anti-CD8 Qdot 655 (custom), anti-CD14 Pacific Blue (BD Biosciences), anti-CD16 Pacific Blue (BD Biosciences), and anti-CD19 Pacific Blue (Invitrogen). Abs for intracellular staining included anti-CD3 Qdot 585 (custom), anti-granzyme B Texas Red PE (BD Pharmingen), anti-IFN-γ Alexa Fluor 700 (BD Pharmingen), and anti-TNF-α PE Cy7 (BD Biosciences). Custom conjugations to Quantum (Q) dot nanocrystals were performed in our laboratory as previously described (25), with reagents purchased from Invitrogen. Anti-human perforin Abs were purchased from Tepnel (clone D48) and BD Biosciences (clone δG9). Additional Abs used for microscopy experiments are listed in the microscopy section.

Cryopreserved PBMC were thawed and then rested overnight at 37°C/5% CO2 in complete medium (RPMI 1640; Mediatech) supplemented with 10% FBS, 1% l-glutamine (Mediatech), and 1% penicillin-streptomycin (Lonza), sterile filtered) at a concentration of 2 × 106 cells/ml of medium in 12-well plates. The next day the cells were washed with complete medium and resuspended at a concentration of 1 × 106 cells/ml with costimulatory Abs (anti-CD28 and anti-CD48d; 1 μg/ml final concentration; BD Biosciences) in the presence of monensin (0.7 μg/ml final concentration; BD Biosciences) and brefeldin A (1 μg/ml final concentration; Sigma-Aldrich). Anti-CD107a was always added at the start of all stimulation periods, as described previously (26). As a negative control, 5 μl of DMSO was added to the cells, an equivalent concentration compared with the peptide stimulus. Staphylococcal enterotoxin B served as the positive control (1 μg/ml final concentration; Sigma-Aldrich). Peptide stimulations were performed at a final concentration of 2 μM. Stimulation tubes were incubated at 37°C/5% CO2 for 6 h. During the time course experiment, stimulation tubes prepared in parallel were incubated for 1, 2, 4, 6, 8, and 12 h before being stained with Abs.

At the end of the stimulation periods, cells were washed once with PBS before being stained for viability with violet or aqua amine-reactive viability dye (Invitrogen) for 10 min in the dark at room temperature. A mixture of Abs was then added to the cells to stain for surface markers for an additional 20 min. The cells were washed with PBS containing 1% BSA (Fisher Scientific) and 0.1% sodium azide (Fisher Scientific) and permeabilized using a Cytofix/Cytoperm kit (BD Biosciences) according to the manufacturer’s instructions. A mixture of Abs against intracellular markers was then added to the cells and allowed to incubate for 1 h in the dark at room temperature. The cells were then washed once with Perm Wash buffer (BD Biosciences) and fixed in PBS containing 1% paraformaldehyde (Sigma-Aldrich). Fixed cells were stored in the dark at 4°C until the time of collection.

HLA-B7/TM10 tetramers were produced according to standard procedures (27). MHC class I tetramer staining was performed in calcium-free PBS for 15 min on ice and in the dark. The cells were then stained for surface and intracellular markers as described above.

For each specimen, between 500,000 and 1,000,000 total events were acquired on a modified flow cytometer (LSRII; BD Immunocytometry Systems) equipped for the detection of 18 fluorescent parameters. Ab capture beads (BD Biosciences) were used to prepare individual compensation tubes for each Ab used in the experiment. Data analysis was performed using FlowJo version 8.7 (Tree Star). Reported data have been corrected for background.

Purified CD8+ T cells were incubated at 37°C in methionine-free medium (Mediatech) for 1 h and then pulsed for 30 min with [35S]methionine (Environmental Health and Radiation Safety, University of Pennsylvania). After washing twice with methionine-free medium, the cells were divided and incubated with control medium or stimulated with PMA (50 ng/ml; Sigma-Aldrich) and ionomycin (1 μM; Sigma-Aldrich) for 4 h. Subsequent immunoprecipitation was performed as described previously (28).

One × 108 YTS cells, either resting or activated via conjugation to KT86 target cells at a 2:1 E:T ratio, were lysed and then subjected to centrifugation at 1000 × g to eliminate the nuclei. Following removal of the nuclei, the remaining lysate was subjected to centrifugation at 18,000 × g to pellet the lytic granules. The pelleted lytic granules and the cytoplasmic fraction were isolated and resuspended in PBS containing protease inhibitors and phosphatase inhibitors, then separated using electrophoresis on a 4–12% Bis-Tris density gradient gel (Invitrogen), and then transferred onto a polyvinylidene difluoride membrane. After blocking with 3% BSA and 140 mM NaCl in TBS, the membrane was incubated with the D48 anti-perforin mAb. Bound Ab was detected with peroxidase-conjugated L chain-specific goat anti-mouse IgG (Jackson ImmunoResearch Laboratories) and an ECL detection system (GE Healthcare). The membrane was stripped in 0.2 M glycine (pH 2.5), 0.05% Tween 20, and 140 mM NaCl in TBS at 50°C for 30 min, blocked with 3% BSA, and reprobed with rabbit β-actin polyclonal Ab clone 20-33 (Sigma- Aldrich), followed by peroxidase-conjugated L chain-specific goat anti-rabbit IgG (Jackson ImmunoResearch Laboratories).

CD8+ T cells were isolated from PBMC by negative selection (Miltenyi Biotec) and then stained with Lysotracker Red (Invitrogen) at a concentration of 166 nM in RPMI 1640 with 10% FCS for 40 min at 37°C, in the dark, washed twice in RPMI 1640 with 10% FCS, as indicated, and allowed to make conjugates with TM10 peptide or irrelevant peptide (HLA-A1-restricted influenza peptide NP44–52, CTELKLSDY)-pulsed C1R-B7 APC at a 2:1 E:T ratio in suspension for either 30 or 240 min. The cell conjugates were then adhered to the poly-l-lysine-coated glass slides and stained as described previously (28). For experiments involving Golgi staining, purified CD8+ T cells were stimulated with either control medium or stimulated with PMA (50 ng/ml; Sigma-Aldrich) and ionomycin (1 μM; Sigma-Aldrich) for 4 h before being stained with anti-Golgi marker Ab (Abcam). For experiments involving HLA staining, the conjugates were stained as described earlier (29), with certain modifications. In brief, conjugates were allowed to adhere on a glass slide as described previously (28) and then blocked with 5% heat-inactivated goat serum in PBS for 20 min at room temperature. Anti-HLA- A/B/C FITC Ab (BD Pharmingen) was diluted in ice-cold PBS containing 2% FCS and 0.1% sodium azide and conjugates were stained in the dark at 4°C for 45 min. Then the slides were rinsed in PBS containing 2% FCS and the cell conjugates were fixed and permeabilized for staining with anti-perforin Abs.

Individual CD8+ T cells or conjugated cells were visualized by using a spinning disc confocal microscope (IX81 DSU; Olympus). Detection settings were adjusted so that a control-stained sample was uniformly negative and fluorescence of experimentally stained samples was not saturating or bleeding through to other channels. The effector and target cells were distinguished by perforin staining. The area of accumulation of fluorescence in the effector cell was measured using an Improvision Volocity software package (PerkinElmer). A threshold of ≥40% of the mean fluorescence intensity of that fluorophore in the cell was used to determine the area occupied. Colocalization between two fluorophores was determined by measuring the area occupied by two different fluorophores. The percent colocalization was then calculated by dividing this colocalized area by the area that is occupied by either of the two fluorophores.

To define the immunological synapse, we measured the percent HLA accumulation at the CTL-target cell interface. The total amount of HLA on the target cell was measured by multiplying the area stained by the HLA-A/B/C Ab (detectable over background) with the mean fluorescence intensity (MFI)4 of the identified region. The effector and target cell regions forming the CTL-target cell interface were selected and measured separately in addition to the total measurement for each conjugate. The percent HLA accumulation (area × intensity) within the interface of each effector-target cell conjugate was then calculated using the formula: percent accumulation = [(synaptic HLA area × synaptic HLA MFI)/(total conjugate HLA area × total conjugate HLA MFI)] × 100. Where specified, the shortest distance of the centroid of the total region of perforin fluorescence and HLA-A/B/C fluorescence at the synapse was measured. All measurements were defined and performed using Improvision Volocity (PerkinElmer).

Targets were prepared from the EBV-transformed cell line C1R, genetically engineered to express HLA-B7 (a gift from Dr. J. Frelinger, University of North Carolina, Chapel Hill, NC). C1R cells expressing HLA-A2 (a gift from Dr. D. Price, Cardiff University, Cardiff, U.K.) were used as a negative control. The cells were incubated with either the B7-restricted peptide TM10 or an irrelevant peptide and (influenza A NP44–52, CTELKLSDY) for 1 h, washed with complete medium, and then labeled with Na51CrO4 for 1 h. Cells were washed three times with RPMI 1640 (Mediatech) before being aliquoted into a 96-well plate (10,000 cells/well). Unlabeled target cells were used as cold targets at the start of the experiment. Effector cells were isolated from donor PBMC through negative selection of CD8+ T cells (Miltenyi Biotec) and were plated according to the various E:T ratios. Where specified, cycloheximide (CHX; 10 μg/ml) was added to the effectors at the time of plating. After 4 h, 51Cr-labeled target cells were added to all of the wells (10,000 cells/well). All test conditions were performed in triplicate and the experiment was repeated four times. Harvested supernatants were evaluated for the presence of 51Cr using a Top Count scintillation counter (Beckman Coulter). The specific lysis was calculated as follows: percent specific lysis = [(mean of the test wells) − (mean of the spontaneous release wells)/(mean of maximal release wells) − (mean of the spontaneous release wells)] × 100%.

Canvas software, version 10.4.9 (ACD Systems) was used to assemble most of the figures. Labels and boxes were added to raw data images in Canvas.

Microscopy.

Significant differences in areas of colocalization, denoted by an asterisk, were calculated using a two-tailed t test (95% confidence interval). For fixed cell microscopy, the minimum number of cells evaluated in a given experiment was determined using a sample size calculation with α and β error levels of 1%. Assumptions were based upon data evaluating colocalization of lytic granule contents published by others (30) and our own preliminary data. For all statistical analyses, differences between cell types or conditions were determined using a two-tailed Student’s t test and were considered significant if p < 0.05.

Chromium release assays.

The data set does not violate the normality assumption. Four independent experiments were conducted to assess the effect of CHX on specific lysis. Since variability in CD8 T cell isolation created unique E:T ratios for each experiment, the relative change in observed percent specific lysis as a result of CHX addition was calculated for every experiment, averaged, and evaluated using a t test with t = 32.80, 3 df. We used repeated measures ANOVA to take into account the “matched” structure of the experiments to be able to look at all of the pairwise differences between the conditions. Since we only focused on the two pairwise comparisons with and without inhibitor, there was no need to adjust the values for multiple comparisons (Tukey-Kramer).

Rapid perforin up-regulation conceivably provides a mechanism by which an Ag-specific CD8+ T cell may retain its cytotoxic capabilities following the exocytosis of preformed secretory granules. To this end, we identified a normal human subject whose CD8+ T cells exhibit a high level of cytotoxicity against CMV TM10 peptide-loaded target cells directly ex vivo in a standard chromium release assay (Fig. 1,A). To address whether newly produced proteins played a role in this killing response, we performed a modified chromium release assay: purified CD8+ T cells were added to peptide-loaded unlabeled target cells for 4 h in the presence or absence of the protein synthesis inhibitor CHX. Following this initial incubation intended to expend preformed granules, chromium-labeled peptide-loaded C1R-B7 cells were added to the effectors and incubated for an additional 4 h. As shown in Fig. 1,B, specific lysis of 40.8% was observed at an initial input E:T ratio of 12.5:1 (effectively a ratio of 2.5:1 when factoring that ∼20% of total donor CD8+ T cells are TM10 specific; see Fig. 2 A) in the absence of CHX, whereas no appreciable lysis was observed against HLA-mismatched targets or irrelevant peptide. Importantly, in the presence of CHX, a marked reduction in the killing ability of TM10-specific CD8+ was observed at all E:T ratios (mean reduction = 0.47 (47%); 95% confidence interval = 0.42–0.51, p < 0.001; t test with df = 3). Four independent experiments yielded similar results (TM10 vs TM10 plus CHX, p = 0.013; paired t test, t = 3.79, df = 5). Thus, de novo protein production generated by Ag-specific CD8+ T cells significantly contributes to continual cytotoxic ability.

FIGURE 1.

New protein synthesis is required for sustained cytotoxicity. A, Purified CD8+ T cells exhibit pronounced ex vivo cytotoxicity. Effector cells were mixed with target cells at a ratio of 15:1. B, Purified CD8+ T cells were added to unlabeled peptide-loaded C1R-B7 cells to flush out preformed perforin from the effector cells. After a 4-h incubation, 51Cr-labeled peptide-loaded C1R-B7 cells were added to the wells and allowed to incubate for an additional 4 h in the presence or absence of the protein synthesis inhibitor CHX to assess the ability of new perforin to sustain cytotoxicity. Shown are representative data of four independent experiments.

FIGURE 1.

New protein synthesis is required for sustained cytotoxicity. A, Purified CD8+ T cells exhibit pronounced ex vivo cytotoxicity. Effector cells were mixed with target cells at a ratio of 15:1. B, Purified CD8+ T cells were added to unlabeled peptide-loaded C1R-B7 cells to flush out preformed perforin from the effector cells. After a 4-h incubation, 51Cr-labeled peptide-loaded C1R-B7 cells were added to the wells and allowed to incubate for an additional 4 h in the presence or absence of the protein synthesis inhibitor CHX to assess the ability of new perforin to sustain cytotoxicity. Shown are representative data of four independent experiments.

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FIGURE 2.

Activated CD8+ T cells rapidly up-regulate perforin. A, Baseline levels of stored perforin (left panels) were determined by staining donor PBMC with HLA-B7/TM10 tetramers. Perforin up-regulation after 1, 2, 4, and 6 h of stimulation with the CMV TM10 peptide was quantified as a function of IFN-γ production (right panels). The red number in each plot represents the proportion of IFN-γ-producing CD8+ T cells that also produce perforin, whereas the black numbers reflect the proportion of total CD8+ T cells that up-regulate perforin. B, Twelve-hour time course experiment, only perforin up-regulation is now measured against degranulation. C, Histograms charting perforin MFI as a function of CD107a expression (left) and IFN-γ production (right). Baseline perforin (dotted line) represents the MFI of perforin stored in the tetramer+ cells at baseline. Events shown are gated upon CD3+CD8+ small lymphocytes, with nonviable cells excluded. D, 35S autoradiogram indicates that the perforin precipitated from PMA-ionomycin (P/I) activated ex vivo CD8+ T cells by the D48 Ab was synthesized de novo. Top, Silver-stained gel containing immunoprecipitates of CTL lysates using IgG, D48, or δG9. Bottom, Autoradiogram of the silver-stained gel. MMM, Molecular mass marker, D48, anti-perforin clone D48; δG9, anti-perforin clone δG9; SEB, staphylococcal enterotoxin B.

FIGURE 2.

Activated CD8+ T cells rapidly up-regulate perforin. A, Baseline levels of stored perforin (left panels) were determined by staining donor PBMC with HLA-B7/TM10 tetramers. Perforin up-regulation after 1, 2, 4, and 6 h of stimulation with the CMV TM10 peptide was quantified as a function of IFN-γ production (right panels). The red number in each plot represents the proportion of IFN-γ-producing CD8+ T cells that also produce perforin, whereas the black numbers reflect the proportion of total CD8+ T cells that up-regulate perforin. B, Twelve-hour time course experiment, only perforin up-regulation is now measured against degranulation. C, Histograms charting perforin MFI as a function of CD107a expression (left) and IFN-γ production (right). Baseline perforin (dotted line) represents the MFI of perforin stored in the tetramer+ cells at baseline. Events shown are gated upon CD3+CD8+ small lymphocytes, with nonviable cells excluded. D, 35S autoradiogram indicates that the perforin precipitated from PMA-ionomycin (P/I) activated ex vivo CD8+ T cells by the D48 Ab was synthesized de novo. Top, Silver-stained gel containing immunoprecipitates of CTL lysates using IgG, D48, or δG9. Bottom, Autoradiogram of the silver-stained gel. MMM, Molecular mass marker, D48, anti-perforin clone D48; δG9, anti-perforin clone δG9; SEB, staphylococcal enterotoxin B.

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To verify that perforin up-regulation may be a mechanism to explain the sustained cytotoxicity we observed in the CTL assay, we monitored the kinetics of perforin up-regulation by flow cytometry relative to degranulation (CD107a expression), granzyme B up-regulation, and cytokine production (IFN-γ, TNF-α). At baseline, 19.3% of the donor CD8+ T cells are bound by the HLA-B7 tetrameric complex containing the CMV peptide TM10 (Fig. 2,A), the majority of which already possess perforin. IFN-γ production was first detected 2 h following stimulation with TM10 peptide. Concomitant with IFN-γ, perforin up-regulation was detected within some activated cells (20.5% of IFN-γ-producing cells) by 2 h (Fig. 2,A, top row). By 6 h, 53.4% of IFN-γ-producing cells were perforin positive. A similar rate of perforin up-regulation was observed in TNF-α-producing cells (data not shown). Interestingly, a second perforin Ab, the commonly used δG9 clone, was unable to detect this early appearance of perforin (Fig. 2 A, bottom row), likely due to the specificity of this Ab for a pH-sensitive motif in granule-associated perforin (24). These results indicate that the kinetics of perforin up-regulation is rapid, coinciding with that of cytokine production.

To demonstrate that we were measuring newly produced perforin, rather than residual unreleased perforin within responding cells, we costained the cells with anti-CD107a to assess degranulation (26). If activated cells were only carrying preformed perforin, then the frequency of CD107a+perforin+ cells should decrease with the exocytosis of cytolytic granules. As demonstrated in Fig. 2,B, CD107a+ cells 1 h poststimulation possess little perforin, indicating perforin loss due to degranulation. Over time, however, the proportion of CD107a+CD8+ T cells that harbor perforin increases, peaking at 53.8% after 8 h, indicating that the responding CD8+ T cells are indeed up-regulating perforin production. The kinetics of granzyme B up-regulation mirrors that of perforin (data not shown). A similar pattern was observed when the cells were stimulated with staphylococcal enterotoxin B and no response was detected with the negative control (Fig. 2,B). Again, the δG9 Ab failed to detect any perforin up-regulation (data not shown). Over the 8-h stimulation period, perforin increasingly accumulates in the CD107a+ and IFN-γ+CD8+ T cell subsets, but does not quite return to baseline levels (Fig. 2 C). This likely reflects an equilibrium between the active production of new perforin and the continued release of perforin, such that a complete return to baseline levels of perforin is only achieved at the conclusion of the immune response.

We also performed a [35S]methionine-labeling experiment on Ag-activated CD8+ T cells to identify whether the perforin recognized by the D48 Ab was newly synthesized after activation (Fig. 2 D). Immunoprecipitation of perforin from activated CD8+ T cells with the D48 Ab yielded a positive signal, but only after T cell activation. In contrast, immunoprecipitation with δG9 resulted in a slight signal before, but not after, activation.

Thus, primary activated human CD8+ T cells rapidly up-regulate perforin de novo, concomitant to cytokine production and degranulation.

To isolate the contribution of new perforin synthesis on sustained cytotoxicity, we used confocal microscopy to monitor the development of new perforin among activated CD8+ T cells. First, we assessed the degree of colocalization between newly up-regulated perforin, detected by the D48 Ab, and granule-associated perforin, stained by the δG9 Ab, in both resting and activated CD8+ T cells purified by negative selection. As shown in Fig. 3,A (top row), both the D48 (blue) and δG9 (green) Abs simultaneously label perforin in resting CD8+ T cells. Moreover, their staining patterns colocalize uniformly with that of Lysotracker Red, which labels the lysosomal granule compartment, indicating that each Ab can recognize granule-associated perforin. After 4 h of stimulation with PMA/ionomycin (Fig. 3,A, second row), however, the proportion of D48-labeled perforin (blue) that colocalizes with that of δG9 (green) decreases from 70% in resting cells to 20% in activated cells (Fig. 3,A, bar graph; D48/δG9). Furthermore, we observed a statistically significant drop in colocalization between D48 perforin (blue) and Lysotracker (red) (Fig. 3,A, bar graph; D48/L 12% in activated cells), whereas the colocalization between δG9 perforin (green) and Lysotracker (red) did not appreciably change (Fig. 3 A, bar graph; δG9/L 25% in resting vs 40% in activated cells). Thus, newly up-regulated perforin can readily be detectable intracellularly and distinct from secretory lysosomes.

FIGURE 3.

New perforin rapidly appears in the Golgi of activated CD8+ T cells. A, Confocal microscopy images illustrating that both the D48 and δG9 Abs recognize the same levels of perforin in resting CD8+ T cells (top row), but not after 4 h of PMA/ionomycin stimulation (second row). Up-regulated perforin in activated CD8+ T cells (blue) resides mostly beyond Lysotracker-labeled granules (red). B, D48-labeled perforin (blue) colocalizes significantly with the Golgi apparatus (orange) of PMA/ionomycin-activated CD8+ T cells, whereas δG9-stained perforin (green) does not. Statistical analysis of the microscopy images: mean area of the cell stained by Abs for each parameter (left panel) and degree of overlap between two Abs (right panel). Combinations represent areas stained by both Abs. For example, G/D48 signifies the proportion of the total Golgi area that is also stained by the D48 Ab. Data generated by analyzing 20 cells from two independent experiments using two different donors. Asterisks denote significant differences (p < 0.05, two-tailed t test) between the area quantified in resting (gray) and activated (black) CD8+ T cells. D48, Anti-perforin clone D48; δG9, anti-perforin clone δG9; G, Golgi; DIC, differential interference contrast.

FIGURE 3.

New perforin rapidly appears in the Golgi of activated CD8+ T cells. A, Confocal microscopy images illustrating that both the D48 and δG9 Abs recognize the same levels of perforin in resting CD8+ T cells (top row), but not after 4 h of PMA/ionomycin stimulation (second row). Up-regulated perforin in activated CD8+ T cells (blue) resides mostly beyond Lysotracker-labeled granules (red). B, D48-labeled perforin (blue) colocalizes significantly with the Golgi apparatus (orange) of PMA/ionomycin-activated CD8+ T cells, whereas δG9-stained perforin (green) does not. Statistical analysis of the microscopy images: mean area of the cell stained by Abs for each parameter (left panel) and degree of overlap between two Abs (right panel). Combinations represent areas stained by both Abs. For example, G/D48 signifies the proportion of the total Golgi area that is also stained by the D48 Ab. Data generated by analyzing 20 cells from two independent experiments using two different donors. Asterisks denote significant differences (p < 0.05, two-tailed t test) between the area quantified in resting (gray) and activated (black) CD8+ T cells. D48, Anti-perforin clone D48; δG9, anti-perforin clone δG9; G, Golgi; DIC, differential interference contrast.

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To further confirm that the perforin labeled by the D48 Ab is produced de novo, we stained the CD8+ T cells with both perforin Abs and an Ab against a 58-kDa Golgi marker protein. As shown in Fig. 3,B, resting CD8+ T cells did not demonstrate a difference in perforin localization detected by either Ab (Fig. 3,B, D48 and δG9, top row). After stimulation with PMA/ionomycin, a large amount of perforin recognized only by the D48 Ab localized within the Golgi (Fig. 3,B, bottom row, D48 plus Golgi), increasing from 15% in resting cells to 47% in activated CD8+ T cells (Fig. 3,B, right bar graph, percent Golgi area also stained by D48, G/D48, p < 0.05; t test). There was no appreciable increase in colocalization between the percent Golgi and the δG9-stained perforin (Fig. 3 B, right bar graph). Similar results were observed in PMA/ionomycin-activated CD8+ T cells from two additional donors (data not shown). Thus, newly produced perforin can be visualized within the Golgi apparatus of activated CD8+ T cells.

To monitor the egress of new perforin from the Golgi, we stained resting and PMA/ionomycin-activated CD8+ T cells with both perforin Abs, as well as an Ab to Rab7 (Fig. 4,A). The latter is a GTPase that characteristically resides on late endosomes and is an important regulator of late endocytic membrane traffic (31). Once again, stimulation with PMA/ionomycin increased the amount of new perforin in the activated cells relative to the resting cells (Fig. 4,A, D48 blue; B, left panel, resting = 1.75 μm2, activated = 4.2 μm2). The colocalization between δG9-stained perforin and Rab7 increased as a result of PMA/ionomycin stimulation (Fig. 4,A, green plus orange; B, right panel, resting = 20%, activated = 55%), likely signifying the formation of secretory lysosomes. In contrast, there was no difference in colocalization between D48-stained perforin (blue) and Rab7 (orange) upon stimulation (Fig. 4,A, blue and orange, and B). Along with the fact that new perforin, as detected by the D48 Ab, does not, for the most part, colocalize with δG9-stained perforin after activation (Fig. 4 B), these data suggest that new perforin is not entirely destined for late endosomal compartments in recently activated cells.

FIGURE 4.

New perforin is not exclusively destined for cytolytic secretory lysosomes. A, Confocal microscopy images of resting CD8+ T cells (top row) and those stimulated with PMA/ionomycin for 4 h (bottom row) stained for Rab7 (orange), perforin (clone D48; blue), and granule-associated perforin (clone δG9; green). B, Left panel, Mean area of the cell stained by each Ab. Combinations represent areas stained by both Abs. Right panel, Degree of overlap between two Abs. For example, R7/D48 signifies the proportion of the total Rab7 area that is also stained by the D48 Ab. Data generated by analyzing 10 cells. Asterisks denote significant differences (p < 0.05, two-tailed t test) between the area quantified in resting (gray) and activated (black) CD8+ T cells. D48, Anti-perforin clone D48; δG9, anti-perforin clone δG9; R7, Rab7. C, Western blot analysis of the perforin content in the lytic granule (Gran) and cytoplasmic (Cyto) fractions of resting (NS) and activated (Stim) YTS NK cells using the D48 Ab. DIC, Differential interference contrast.

FIGURE 4.

New perforin is not exclusively destined for cytolytic secretory lysosomes. A, Confocal microscopy images of resting CD8+ T cells (top row) and those stimulated with PMA/ionomycin for 4 h (bottom row) stained for Rab7 (orange), perforin (clone D48; blue), and granule-associated perforin (clone δG9; green). B, Left panel, Mean area of the cell stained by each Ab. Combinations represent areas stained by both Abs. Right panel, Degree of overlap between two Abs. For example, R7/D48 signifies the proportion of the total Rab7 area that is also stained by the D48 Ab. Data generated by analyzing 10 cells. Asterisks denote significant differences (p < 0.05, two-tailed t test) between the area quantified in resting (gray) and activated (black) CD8+ T cells. D48, Anti-perforin clone D48; δG9, anti-perforin clone δG9; R7, Rab7. C, Western blot analysis of the perforin content in the lytic granule (Gran) and cytoplasmic (Cyto) fractions of resting (NS) and activated (Stim) YTS NK cells using the D48 Ab. DIC, Differential interference contrast.

Close modal

To confirm these observations, we analyzed the perforin content of lytic granules and the cytoplasmic fraction of activated NK cells. We used YTS NK cells, which like ex vivo human NK cells constitutively express high levels of perforin, for easier detection of perforin by Western blot. As can be seen in Fig. 4 C, perforin is readily detected in the lytic granules both before and after stimulation; however, following conjugation with KT86 target cells, there is a substantial accumulation of new perforin in the cytoplasmic fraction (increase in band intensity from 0.3 to 0.6 by densitometry).

The fact that new perforin does not immediately localize with late endosomes suggests that it is not intimately involved in sustaining cytotoxicity. However, a precedent for an alternative perforin secretion pathway exists (23). We therefore directly visualized the fate of newly produced perforin (D48 labeled) in activated Ag-specific CD8+ T cells incubated with HLA-B7-transfected C1R APC loaded with either the cognate TM10 peptide or an irrelevant peptide. After 30 and 240 min of incubation, the cell conjugates were stained with both perforin Abs (δG9 and D48 clones) to permit the visualization of granule-associated (δG9 and D48) and newly produced (D48) perforin, as well as Lysotracker. When the APC were pulsed with irrelevant peptide and mixed with CD8+ T cells for 30 min (Fig. 5,A), the granules appeared distal to the CTL-target interface. When exposed to APC primed with TM10 peptide (Fig. 5,B), the perforin-containing granules reoriented toward the interface. There was no change in perforin content stained by either the D48 or δG9 Ab at 30 min (Fig. 5 C).

FIGURE 5.

Cytotoxic secretory lysosomes polarize toward the immunological synapse after 30 min of conjugation with cognate target cells. CD8+ T cells, purified from donor PBMC by negative selection, were allowed to form conjugates for 30 min with HLA-B7-C1R APC presenting nonspecific flu peptide (A) or peptide TM10 (B) and then stained for perforin (clone D48; blue), granule-associated perforin (clone δG9; green), and with Lysotracker Red (red). All images are oriented such that the T cell is at the top and the target cell is at the bottom. C, Left panel, Mean area of the CD8+ T cell stained by each Ab. Combinations represent areas stained by both Abs. Right panel, Degree of colocalization between two Ab-stained areas. For example, D48/δG9 represents the proportion of the total D48 area that is also stained by the δG9 Ab. D, To define the immunological synapse, the same conjugates were stained in parallel for HLA-A/B/C molecules (red). All images are oriented such that the CD8+ T cell is at the top and the target cell is at the bottom. E, Percentage of total HLA-A/B/C accumulated at the CTL-target cell interface. All data were generated by analyzing 10 conjugates. Asterisks denote significant differences (p < 0.05, two-tailed t test). D48, Anti-perforin clone D48; δG9, anti-perforin clone δG9; L, Lysotracker dye; DIC, differential interference contrast.

FIGURE 5.

Cytotoxic secretory lysosomes polarize toward the immunological synapse after 30 min of conjugation with cognate target cells. CD8+ T cells, purified from donor PBMC by negative selection, were allowed to form conjugates for 30 min with HLA-B7-C1R APC presenting nonspecific flu peptide (A) or peptide TM10 (B) and then stained for perforin (clone D48; blue), granule-associated perforin (clone δG9; green), and with Lysotracker Red (red). All images are oriented such that the T cell is at the top and the target cell is at the bottom. C, Left panel, Mean area of the CD8+ T cell stained by each Ab. Combinations represent areas stained by both Abs. Right panel, Degree of colocalization between two Ab-stained areas. For example, D48/δG9 represents the proportion of the total D48 area that is also stained by the δG9 Ab. D, To define the immunological synapse, the same conjugates were stained in parallel for HLA-A/B/C molecules (red). All images are oriented such that the CD8+ T cell is at the top and the target cell is at the bottom. E, Percentage of total HLA-A/B/C accumulated at the CTL-target cell interface. All data were generated by analyzing 10 conjugates. Asterisks denote significant differences (p < 0.05, two-tailed t test). D48, Anti-perforin clone D48; δG9, anti-perforin clone δG9; L, Lysotracker dye; DIC, differential interference contrast.

Close modal

After 240 min, the amount of total D48 Ab-stained perforin increased in the TM10-APC/CD8+ T cell conjugates, but not in the control samples (Fig. 6,A). The residual perforin contained in the cytolytic lysosome compartment, as stained by the δG9 Ab (green) and Lysotracker (red), occupied a very specific area at the center of the interface (Fig. 6,A, δG9 plus Lysotracker, bottom row). Perforin stained by the D48 Ab (blue), also appeared at the interface, but occupied a substantially larger area of the cell (Fig. 6,A, overlay, bottom row). The proportion of D48 staining that colocalized with Lysotracker decreased from ∼39% in resting cells to ∼26% in activated cells (Fig. 6 B, D48/L).

FIGURE 6.

New perforin accumulates at the immunological synapse largely independent of secretory lysosomes. A, CD8+ T cells, purified from donor PBMC by negative selection, were allowed to form conjugates for 240 min with HLA-B7-C1R APC presenting peptide TM10 or nonspecific (control) flu peptide and then stained for perforin (clone D48; blue), granule-associated perforin (clone δG9; green), and with Lysotracker Red (red). All images are oriented such that the T cell is at the top and the target cell is at the bottom. B, Left panel, Mean area of the TM10/C1R (▪)- or irrelevant peptide/C1R ()-conjugated CD8+ T cells stained by each parameter. Combinations represent areas stained by both Abs. Right panel, Degree of colocalization between two Ab-stained areas. For example, D48/δG9 represents the proportion of the total D48 area that is also stained by the δG9 Ab. C, To define the immunological synapse, conjugates between purified donor CD8+ T cells and TM10 or nonspecific (control) flu peptide-loaded HLA-B7-C1R cells were stained in parallel for HLA-A/B/C molecules (red). D, Percentage of total HLA-A/B/C that accumulated at the CTL-target cell interface. E, The shortest distance between the centroid of the D48-defined perforin region and HLA-A/B/C at the effector-target cell interface was measured for both TM10- and irrelevant peptide- loaded targets after 30 and 240 min of conjugation. Mean values ± SD are shown. All data were generated by analyzing ≥10 conjugates. Asterisks denote significant differences (p < 0.05, two-tailed t test) between the area quantified in nonspecific conjugates () and TM10-specific (▪) conjugates. D48, Anti-perforin clone D48; δG9, anti-perforin clone δG9; L, Lysotracker dye: DIC, differential interference contrast.

FIGURE 6.

New perforin accumulates at the immunological synapse largely independent of secretory lysosomes. A, CD8+ T cells, purified from donor PBMC by negative selection, were allowed to form conjugates for 240 min with HLA-B7-C1R APC presenting peptide TM10 or nonspecific (control) flu peptide and then stained for perforin (clone D48; blue), granule-associated perforin (clone δG9; green), and with Lysotracker Red (red). All images are oriented such that the T cell is at the top and the target cell is at the bottom. B, Left panel, Mean area of the TM10/C1R (▪)- or irrelevant peptide/C1R ()-conjugated CD8+ T cells stained by each parameter. Combinations represent areas stained by both Abs. Right panel, Degree of colocalization between two Ab-stained areas. For example, D48/δG9 represents the proportion of the total D48 area that is also stained by the δG9 Ab. C, To define the immunological synapse, conjugates between purified donor CD8+ T cells and TM10 or nonspecific (control) flu peptide-loaded HLA-B7-C1R cells were stained in parallel for HLA-A/B/C molecules (red). D, Percentage of total HLA-A/B/C that accumulated at the CTL-target cell interface. E, The shortest distance between the centroid of the D48-defined perforin region and HLA-A/B/C at the effector-target cell interface was measured for both TM10- and irrelevant peptide- loaded targets after 30 and 240 min of conjugation. Mean values ± SD are shown. All data were generated by analyzing ≥10 conjugates. Asterisks denote significant differences (p < 0.05, two-tailed t test) between the area quantified in nonspecific conjugates () and TM10-specific (▪) conjugates. D48, Anti-perforin clone D48; δG9, anti-perforin clone δG9; L, Lysotracker dye: DIC, differential interference contrast.

Close modal

To define the CTL-target cell interface as the immunological synapse, we assessed the degree of MHC class I clustering at the contact zone between the CTL and target cells. After 30 min of incubation, we observed a significant amount of MHC class I accumulation at the interface between the CTL and TM10-loaded target cells (Fig. 5,D, red, bottom row) as compared with that of target cells loaded with irrelevant peptide (Fig. 5,D, top row; E, TM10 68% of HLA vs control peptide 34%; p = 0.0254, two-tailed t test). By 240 min of conjugation formation, the degree of HLA clustering at the interface remained pronounced (Fig. 6,C, bottom row, red; D, 68% for 30 min vs 67% for 240-min conjugates, p = 0.0049, two-tailed t test). The D48-labeled perforin (blue) and δG9-labeled perforin (green) accumulated at the immunological synapse (overlay of Figs. 5,D and 6 C) only when the CD8+ T cells received an activation signal by TM10-pulsed APC.

We next calculated the distance between the D48-labeled perforin centroid and the HLA class I cluster at the CTL-target cell interface to define the polarization of newly synthesized perforin to the immunological synapse. We found this distance to be significantly less between CD8+ T cells and TM10-loaded APC compared with APC-bearing irrelevant peptide at both 30 min (Fig. 6,E, TM10 peptide = 1.66 μm vs irrelevant peptide = 3.68 μm; p = 0.014, two-tailed t test) and 240 min of conjugation (Fig. 6,E, TM10 peptide = 1.49 μm vs irrelevant peptide = 3.66 μm; p = 0.008, two-tailed t test). The former data reflect the migration of stored perforin toward the synapse immediately following activation, as evidenced by the similar localization of the D48- and δG9-labeled perforin (Fig. 5,D, bottom row, D48 plus δG9, and from the parallel analysis in C), whereas the latter data are mostly attributable to the polarization of new perforin (Fig. 6 C, bottom row, D48 (blue) vs δG9 (green), overlay). These data suggest that within activated CD8+ T cells newly produced perforin is transported to the immunological synapse, where it may promote sustained cytotoxic activity.

It has long been appreciated that CD8+ T cells play a pivotal role in the elimination of virally infected cells and that perforin is a key mediator of this process through its distinct ability to enable the entry of apoptosis-inducing granzymes. The mechanism for perforin-mediated Ag-specific killing has previously been attributed to the exocytosis of cytotoxic granules present within the CD8+ T cell. In this study, we redefine this mechanism, demonstrating that virus-specific CD8+ T cells rapidly up-regulate perforin after activation and then target the protein directly to the immunological synapse. This continual production and targeted release of perforin after stimulation may allow the CD8+ T cell to recognize and kill additional targets after the initial release and depletion of the cell’s complement of preformed cytotoxic granules. Additionally, our results demonstrate that multiple forms of perforin are present within resting and activated CD8+ T cells, which are not necessarily constrained within cytolytic lysosomes. This indicates that many aspects of perforin protein regulation, expression, trafficking, structure, and mechanism of action remain to be elucidated.

Previous studies have suggested that perforin expression by CD8+ T cells requires entry into the cell cycle and subsequent proliferation (20, 21, 22). If this were so, CD8+ T cells would have a substantial period of time after initial target recognition during which they would essentially be disarmed. Although increased perforin mRNA after stimulation has been observed previously, the failure to detect new perforin protein following activation has prompted the conclusion that perforin expression is linked to the proliferative potential of the cell. Contrary to these previous observations, we find that Ag-specific CD8+ T cells can up-regulate perforin protein production in as little as 2 h after TCR stimulation directly ex vivo, concordant with the kinetics of perforin mRNA up-regulation (24). Thus, perforin up-regulation and expression clearly does not require proliferation.

Perhaps most importantly, our results indicate that de novo production of perforin following Ag-specific stimulation is biologically relevant, for when this process is inhibited by the protein-synthesis inhibitor CHX, a dramatic reduction in killing ability results. Because CHX is not solely perforin specific, it is possible that the observed decrease in cytotoxicity is due to the inhibition of additional proteins beyond perforin, such as Fas ligand, granzymes, or TNF-α, that are also involved in mediating target cell apoptosis. Specifically targeting perforin for inhibition in human CD8+ T cells ex vivo is not trivial, but alternative avenues are currently being explored in our laboratory.

Notably, newly formed perforin can also be transported directly to the immunological synapse independent of cytolytic granules upon Ag-specific stimulation. The existence of a secondary pathway of perforin exocytosis has previously been described (23), but was postulated to be a nonspecific process that leads to bystander killing instead of Ag-specific killing. Our results confirm the existence of an alternative perforin exocytosis pathway and indicate that CD8+ T cells can specifically employ this pathway to deliver newly formed perforin to the target cell contact site.

A recent study reported the cooperation between a lysosomal cytotoxic granule and an endosomal exocytic vesicle as a prerequisite for the cytotoxic function of lymphocytes (30). The effector protein hMunc13-4 coordinates the assembly of the two organelles and then primes granule fusion at the CTL-target cell interface. Our results do not contradict, but rather complement, this report. Whereas Menager et al. (30) used the δG9 anti-human perforin Ab to examine the mechanism for cytolytic granule-associated perforin release, our study focused upon the production and release of newly formed perforin. It remains to be determined whether newly formed perforin also transitions through the same exocytosis pathway controlled by hMunc13-4. What other organelles and cofactors mediate the transport of the new perforin directly to the immunological synapse are critically important to define for full understanding of T cell effector function.

We submit, therefore, that cytolytic granules are necessary for the proper storage of perforin and granzymes in resting CTL, making them available for immediate release upon TCR triggering. Upon activation, however, new perforin does not need to progress via the secretory lysosome pathway to be released from the cell. Although some of the new perforin may be allocated to replenish the granules, an appreciable quantity is directly targeted to the immunological synapse to mediate cytotoxicity, thereby providing a means for continual and repeated cytotoxicity. Although secretion of cytolytic molecules has been described previously through the constitutive pathway (23), we were surprised to find that this can occur in a directed manner to participate in targeted cytotoxicity. This phenomenon is not unlike the bidirectional release of cytokines described for activated Th cells, in which distinct trafficking proteins are associated with each secretion pathway (32, 33). Whereas Th cells secrete cytokines and chemokines in multiple directions by different pathways, we propose that two separate trafficking routes are responsible for delivering both new perforin and preformed perforin stored in granules to the site of contact with the target cell.

Perforin requires additional modifications following its egress from the Golgi to achieve its active form, which typically occur in the cytolytic granule. How then does newly produced perforin achieve the necessary conformation required for pore formation if it does not transit via the secretory lysosomal pathway? If newly formed perforin does not achieve an active conformation within the secretory lysosomes, then it could either be modified within the immunological synapse, on the target cell membrane itself, or within an endosomal vesicle inside the target cell. The answer to this question is likely intertwined with the mechanism and location of perforin’s action, which remains controversial.

We thank Dr. Jeff Frelinger and Dr. David Price for generously donating the C1R cells expressing HLA-B7 and A2, respectively. Dr. John Wherry provided the HLA-B7/TM10 peptide tetramer reagent. Linda Monaco-Shawver deserves notable praise for assisting with the chromium release assays, as does Jay Gardner for generating custom Qdot-conjugated Abs. We appreciate the efforts of the Human Immunology Core at the University of Pennsylvania, directed by Dr. James Riley, for furnishing us with donor PBMC samples and the anti-HLA-A/B/C Ab, as well as the Flow Cytometry and Cell Sorting Facility of the Abramson Cancer Center at the University of Pennsylvania for continued technical support. Christie Bell also deserves special recognition for technical assistance in optimizing the detection of perforin up-regulation. We thank Drs. Mario Roederer and Guido Silvestri for critical discussion of the results.

The authors have no financial conflict of interest.

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 the following grants and organizations: National Institutes of Health Grants AI076066 (to M.R.B.), AI067946 (to J.S.O.), and AI079731 (to J.S.O.) and the W. W. Smith Foundation (to M.R.B.).

4

Abbreviations used in this paper: MFI, mean fluorescence intensity; CHX, cycloheximide.

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