Published ahead of print on June 5, 2003, doi:10.1165/rcmb.2002-0314OC
© 2003 American Thoracic Society DOI: 10.1165/rcmb.2002-0314OC Proapoptotic Effects of Parathyroid Hormone-Related Protein in Type II PneumocytesResearch, Anesthesiology, and Medicine Services, VA San Diego Healthcare System, San Diego; and Departments of Anesthesiology and Medicine, University of California San Diego, La Jolla, California Address correspondence to: Randolph H. Hastings, M.D., Ph.D., VA Medical Center (125), 3350 La Jolla Village Dr., San Diego, CA 92161–5085. E-mail: rhhastings{at}ucsd.edu
Parathyroid hormone–related protein (PTHrP) promotes or suppresses apoptosis in various settings depending on cell type and context. PTHrP 1–34 and PTHrP 67–86 are type II cell growth factors with effects on pneumocyte growth and surfactant secretion. This study investigated the effects of 24 h pretreatment with these two peptides on rat type II cell apoptosis after 0.3 J/cm2 ultraviolet-B irradiation. Adherent cells decreased in number by 15 ± 5% and nonadherent cells increased > 5-fold 24 h after ultraviolet irradiation. Cell loss was due predominantly to apoptosis, based on ethidium bromide exclusion, nuclear condensation, and caspase 3 activity. Nuclear condensation increased from 15.6 ± 2.2% of irradiated cells with no treatment to 25.6 ± 4.9 and 22.9 ± 1.8% of cells in ultraviolet/PTHrP 1–34 and ultraviolet/PTHrP 67–86 groups, respectively (P < 0.01), along with a 60% increase in caspase 3 activity. Effects on apoptosis were unaffected by the presence or absence of serum, but were ameliorated by growth to confluence or adherence to fibronectin. PTHrP 1–34 and PTHrP 67–86 augmented inositol phosphate levels, but had minimal effects on cAMP. Thus, PTHrP 1–34 and PTHrP 67–86 sensitize type II cells to apoptosis, possibly by a phospholipase C–dependent mechanism. The effects appear to be regulated by cell–matrix and cell–cell interactions.
Abbreviations: 7-amino-4-methyl coumarin, AMC • aspartyl-glutamyl-valyl-aspartate, DEVD • 3-[3-(Cholamidopropyl)dimethylammonio]-1-proanesulfonate, CHAPS • dimethylsulfoxide, DMSO • ethylene diamine tetraacetic acid, EDTA • ethylene Glycol-bis(beta-aminoethyl-ether)-N,N,N',N'-tetraacetatic acid, EGTA • fetal bovine serum, FBS • fluoromethylketone, fmk • human embryonic kidney, HEK • N-2-Hydroxyethylpiperazine-N'-2-ethanesulfonic acid, HEPES • high powered field, hpf • phosphate-buffered saline, PBS • piperazine-1,4-bis(2-ethanesulfonic acid), PIPES • parathyroid hormone, PTH • parathyroid hormone-related protein, PTHrP • relative fluorescent units, RFU
Parathyroid hormone–related protein (PTHrP) was discovered as a hypercalcemia-inducing factor in squamous cell carcinomas (1). PTHrP 1–34 has structural similarities to parathyroid hormone (PTH) 1–34, binds the same receptor, and mimics all of the effects of PTH in tissues bearing the receptor including hypercalcemia. PTHrP is produced in many normal tissues, where it exerts local effects related to growth, skeletal metabolism, and smooth muscle relaxation (2). The growth-related effects include tissue-specific stimulation or inhibition of apoptosis (3–7). For example, PTHrP 1–141 overexpression protects chondrocytes and pancreatic ß cells against apoptosis (3, 4), and exogenous PTHrP 1–34 protects ß cells (4). In contrast, PTHrP is proapoptotic in mesenchymal cells, kidney epithelial cells, and IEC-6 intestinal epithelial cells (5–7). Mid-molecule and carboxy-terminal peptides of PTHrP are also biologically active and have effects independent of PTHrP 1–34. PTHrP 109–141 stimulates osteoclast retraction, inhibits osteoclast tartrate-resistant acid phosphatase activity, and stimulates cAMP production in osteoblasts (8). PTHrP 67–86 inhibits mitogenesis and stimulates the metastatic potential of human breast carcinoma cells (9). PTHrP is expressed and secreted in the lung by alveolar type II epithelial cells (10). Type II cells also express the PTH/PTHrP receptor (11), which signals through protein kinase A, protein kinase C, and intracellular calcium pathways (12). PTHrP 1–34 stimulates phosphatidylcholine production and secretion in cultured type II cells (11) whereas neutralizing PTHrP antibodies induce proliferation of type II cells in culture, and type II cells in vivo (13). Thus, PTHrP has autocrine effects on type II cell function and inhibits type II cell growth. Recently, we demonstrated that a mid-molecule peptide, PTHrP 67–86, stimulates type II cell phosphatidylcholine secretion (14), similar to the effect of PTHrP 1–34. Type II cells undergo apoptosis in fibrotic lung and after various forms of lung injury (15–18). Apoptosis may play a homeostatic role in repair after lung injury by controlling type II cell growth and may help to restore the normal alveolar architecture by eliminating excess pneumocytes (15). Because PTHrP is an autocrine growth factor for type II cells and regulates apoptosis in other cell types, it could have a role in regulating type II cell death. The overall goal of our study was to determine whether two lung-active PTHrP peptides, PTHrP 1–34 and PTHrP 67–86, alter the sensitivity of type II cells to apoptosis. Ultraviolet radiation has been used to induce type II cell apoptosis by other investigators (18, 19) and was chosen as the stimulus in this project because it caused reproducible degrees of apoptosis in preliminary studies. Ultraviolet can cause apoptosis through death-receptor or mitochondrial-initiated pathways, as can lung injury (20–23). Thus, studies of ultraviolet-induced apoptosis may elucidate pathways common to pneumocyte apoptosis in the setting of acute lung injury. A second goal was to begin investigations into regulatory mechanisms. We studied modulatory influences of confluence, serum, and caspase 3 inhibition. Because PTHrP 1–34 and PTHrP 67–86 have similar effects on type II cell function and were found to have similar effects on apoptosis, a goal of this study was to compare the effects of the two peptides on secondary messenger levels. We also evaluated Akt kinase activation and confluence, important factors in how PTHrP alters sensitivity to apoptosis in mesenchymal cells (5).
General Protocols Type II cell isolation. Type II cells were isolated from healthy 200- to 250-g rats by the method of Dobbs (10, 24). Briefly, the lungs were perfused and lavaged to remove airway inflammatory cells, digested with low concentrations of elastase, minced, filtered through gauze and nylon screen filters, and washed free of protease. Type II cells were isolated from the crude cell preparation by panning on IgG coated plates. Purity was determined from wet mounts of cells stained with Phosphine 3R (Polysciences, Warrington, PA). Cell yield averaged 35 ± 3 x 106 cells/isolation and type II cell purity averaged 74 ± 1% (n = 23 isolations). Cell culture. For nuclear condensation experiments, type II cells were plated at 400,000 cells/well in NUNC Slide-Tek wells with DMEM plus 10% fetal bovine serum (FBS). The epithelial cells were plated in 12-well tissue culture plates (BD Biosciences, Franklin Lakes, NJ) at 3 x 106 cells/well for caspase assays. The pneumocytes were incubated at 37°C in a humidified incubator with 10% CO2 / 90% air for 18 h before experiments. Jurkat cells were cultured in suspension in RPMI 1640 plus 10% FBS. Treatments. Cells were washed three times with phosphate-buffered saline (PBS) and treated with 1 µM PTHrP 1–34 (Bachem, Torrance, CA), 1 µM PTHrP 67–86 (Genemed Synthesis, South San Francisco, CA), or media plus serum alone for 24 h before ultraviolet exposure. Other investigators have used comparable PTHrP concentrations in apoptosis studies (4–6). Micromolar concentrations are high compared with reported effective range for secondary messenger production (12), but may reproduce ambient concentrations in the vicinity of receptors after autocrine release of PTHrP. The PTHrP peptides were dissolved in sterile H2O and the no treatment group contained an equivalent volume of dH2O as given in the PTHrP treatment groups. Ultraviolet irradiation. Type II cells were irradiated with a shortwave ultraviolet transilluminator (Fotodyne, Inc., New Berlin, WI) for 10 min at room temperature in a sterile hood. The energy delivered by the shortwave ultraviolet transilluminator to the cells was calibrated using an ultraviolet detection J-meter (UVP, Upland, CA) under a disk that was plated with cells. The measured energy output was 0.3 J/cm2 at 310 nm. Apoptosis assays were generally performed in media with serum and with cells at 60% confluence, except for certain experiments (see SPECIFIC PROTOCOLS). The cells were then allowed to recover for 24 h at 37°C in a humidified incubator before further processing for apoptotic markers.
Specific Protocols (2) Modulation of PTHrP peptide effects in serum-free media. To investigate whether the effects of PTHrP on apoptosis depend on growth factors in serum, cells were incubated in serum-free media with 0.5% bovine serum albumin during the pretreatment period, the ultraviolet exposure, and the subsequent 24-h recovery period. Caspase 3 activity was used as the measure of apoptosis. (3) Modulation of PTHrP peptide effects by substrate. Caspase 3 activity was assayed in cells plated in media plus serum on fibronectin-coated (4 µg/cm2) plates after ultraviolet exposure and recovery with and without PTHrP peptide pretreatment, as described above. (4) Modulation of PTHrP peptide effects by confluence. Cells were plated at the same density as in the preconfluent experiment and cultured for an extra day to reach confluence. PTHrP treatment, ultraviolet exposure, and recovery followed as described for protocol #1. Caspase 3 assay was used as the measure of apoptosis. (5) Effect of caspase 3 inhibition on ultraviolet-induced apoptosis. In some experiments, cells were pretreated with a specific caspase 3 inhibitor, acetyl-aspartyl -glutamyl -valyl-glutamyl-fluoromethylketone (Ac-DEVD-FMK; Calbiochem, La Jolla, CA). Stock Ac-DEVD-fmk in dimethylsulfoxide (DMSO) was added to the media plus serum to a final concentration of 0.1 mM with 1:1,000 vol:vol DMSO. Control cells were treated with an equal volume of DMSO. Cells were pretreated with PTHrP peptides for 24 h and caspase 3 inhibitor for the final 30 min before ultraviolet exposure. Caspase 3 activity was measured after 24 h recovery. Apoptosis was assessed independently by nuclear condensation measurements. (6) Effect of PTHrP 1–34 and PTHrP 67–86 on secondary messenger levels. Type II cells were plated in 12-well plates and studied 24 h after isolation. Cells were washed with PBS and then stimulated with PTHrP 1–34 or PTHrP 67–86 in serum-free media for 10 min. The positive control was 0.1 mM forskolin for cAMP production and ATP for inositol phosphate stimulation. Secondary messengers were assayed as described below. Duplicate wells were studied for cells from each rat and experimental group.
Measurements Cell death measurements. Conditioned media were removed and cells were washed once with PBS. Adherent cells were treated with 0.12 µM ethidium bromide and 4 µM Hoescht 33342 in media. Ethidium bromide is excluded from viable cells, whereas H33342 is a cell permeant bis-benzimide dye that labels all nuclei. The fraction of dead cells was determined by counting ethidium bromide–positive cells and dividing by the total number of nuclei. At least 150 cells were counted per experimental treatment per animal. Nonadherent cells were evaluated for viability by incubating cells for 2 min with trypan blue. Blue-stained cells were considered nonviable. Cell number and nuclear condensation assessment. Adherent cells were washed with PBS and fixed with 100% methanol. Nuclei were stained with Hoescht 33342. Fluorescent images were taken with a Kodak 290 digital camera system, saved as color TIFF files, and converted to 8-bit grayscale for image analysis. Cell number per 400x high-power field (hpf) and nuclear area were quantitated using NIH Image 1.62 software. Type II cells that were not exposed to ultraviolet nor treated with PTHrP were used to establish a lower threshold for the area of nuclei in nonapoptotic cells. Condensed nuclei were defined as those nuclei whose areas were less than or equal to the fifth percentile for nuclear areas of normal cells. To evaluate nonadherent cells for apoptosis, cytospin preparations were fixed, stained, and examined as described for adherent cells. At least 200 cells were examined per experimental condition and cell isolation, and at least 300 cells were evaluated per control group in establishing the cutoff area. Caspase 3 assay. Adherent cells were washed once in PBS, scraped in 500 µl caspase lysis buffer (50 mM PIPES/KOH, 2 mM EDTA, 0.1% [wt/vol] CHAPS, 1 mM dithiothreitol, 20 µg/ml leupeptin, and 1 µg/ml pepstatin A), pooled with the nonadherent cells from the same wells and lysed through sonication. Lysates were kept at 4°C until time of assay. Protein was measured with the Bio-Rad protein assay kits. Assays were performed in 96-well plates with 20 µg total protein from cell lysates. Caspase 3 substrate, Ac-DEVD-7-amino-4-methyl coumarin (AMC), and inhibitor, acetyl-DEVD-CHO, (Alexis Biochemicals, Carlsbad, CA) were diluted separately to 0.1 mM concentration in caspase assay buffer (50 mM HEPES, 100 mM NaCl, 0.1% [wt/vol] CHAPS, 5 mM dithiothreitol). The caspase inhibitor was used in separate control assays to test the specificity of the reaction. It was incubated with the lysate samples at room temperature for 20 min before substrate. Assays were initiated by heating the plates to 37°C and adding the substrate to the sample wells. Fluorescence of AMC cleaved from the substrate was detected on a 96-well multilabel reader (Perkin Elmer, Boston, MA) at excitation and emission wavelengths of 355 nm and 460 nm, respectively, with a 0.5-s capture window. Fluorescent readings were taken at 30-min intervals, exported to Microsoft Excel, and converted to relative fluorescent units (RFUs) by subtracting the background fluorescence of the reagent blank for each plate to compensate for plate-to-plate variability. Caspase 3 activity was calculated as the slope of the plot of RFU versus time over the initial linear portion of the curve, generally the first 30–60 min. Activities were normalized to the mass of cell protein used in the assay. Immunoblotting. Cells were collected by trypsinization and pelleted by centrifugation at 200 x g for 5 min. The pellets were washed with PBS and then solubilized with cold 50 mM PIPES/KOH, pH 6.5, 0.1% CHAPS, 2 mM EDTA, 1 µM pepstatin, 10 µM leupeptin, and 0.2 mM phenylmethylsulfonyl fluoride. Lysate samples were cleared of debris by centrifugation at 20,000 x g for 15 min at 4°C and heated to 70°C for 10 min in Laemmli sample buffer with 10% ß-mercaptoethanol. Proteins (10 µg cell protein/lane) were separated by electrophoresis on 4–12% SDS polyacrylamide gels (Invitrogen Corp., Carlsbad, CA), and electroblotted onto nitrocellulose membranes (Millipore Corp., Bedford, MA). Membranes were washed with Tris-buffered saline with 0.5% Tween 20 (TBS-T) and blocked for 1 h with 10% milk protein in dH2O at 4°C. Membranes were incubated for 1 h with primary antibody at 4°C, washed extensively with TBS-T, then incubated with horseradish peroxidase–conjugated anti-mouse or goat IgG secondary antibody for 45 min. Bands were visualized by chemiluminescence using the PicoSignal (Pierce Chemical, Rockford, IL) and quantitated by densitometry (Alpha Innotech Corp., San Leandro, CA). Primary antibody to Akt-1, -2, and -3 kinase phosphorlyated at Ser 473, was obtained from Cell Signaling Technologies, Inc. (Beverly, MA), whereas the actin antibody came from Santa Cruz Biotechnology (Santa Cruz, CA). The positive control for phosphorylated Akt was a Jurkat cell lysate. cAMP and inositol phosphate assay. Cells were extracted with 1 mM HCl and cAMP was measured by radioimmunoassay (DELPHI, Perkin and Elmer Biosciences). To measure inositol phosphates, cells were loaded for 24 h with 2µCi [3H]-myo inositol (New England Nuclear Life Science, Boston, MA), then washed and stimulated with the PTHrP peptides or 100 µM ATP. The cells were extracted with 0.1 N HCl/100% methanol (1:1 vol:vol) and spun at 2,000 x g for 2 min. The supernatant was passed over Dowex AG 1-X8 columns and washed twice with water and twice with 60 mM ammonium formate to elute the free myo-inositol. Inositol phosphates were eluted with 1 M ammonium formate in 100 mM formic acid and quantified by scintillation counting.
Data Analysis
Effect of PTHrP 1–34 and PTHrP 67–86 on Type II Cell Death and Apoptosis after Ultraviolet Irradiation Cell loss after ultraviolet exposure. Figure 1 demonstrates the effects of ultraviolet exposure and PTHrP peptide treatment on the retention of adherent type II cells from nine independent cell isolations. Cell densities decreased by 15% with ultraviolet exposure, and treatment with PTHrP 1–34 or PTHrP 67–86 resulted in a loss of an additional 15% in the irradiated cells (P < 0.05 for PTHrP 1–34, P = 0.06 for PTHrP 67–86). Ultraviolet irradiation increased the number of nonadherent cells/well 5-fold compared with controls (P < 0.01, Figure 2), and treatment with PTHrP 1–34 or PTHrP 67–86 augmented the effect of the radiation on inducing cell loss from the Slide-Tek well surface. Irradiation without treatment resulted in 14,600 ± 3,800 nonadherent cells/well compared with 22,500 ± 7,800 and 27,900 ± 5,000 cells/well with PTHrP 1–34 and PTHrP 67–86 treatment, respectively (**P < 0.001 versus control and ultraviolet untreated cells, n = 6–9 cell isolations). Neither PTHrP peptide affected the number of adherent and nonadherent cells in nonirradiated cells. Cell numbers did not differ between PTHrP 1–34 and PTHrP 67–86 treatments.
Effects of PTHrP peptides on ultraviolet-induced cell death. ultraviolet exposure caused a 4- to 6-fold increase in the number of adherent cells that took up ethidium bromide, but the percentage remained  5% (Table 1). Treatment with PTHrP 1–34 or PTHrP 67–86 did not cause a significant increase in the percentage of ethidium bromide–positive cells after ultraviolet exposure. However, the percentage was greater in nonirradiated cells treated with PTHrP 1–34 than in untreated cells or cells treated with PTHrP 67–86.
Effect of PTHrP peptides on ultraviolet-induced apoptosis. The percentage of cells with condensed or fragmented nuclei increased from 5.0 ± 0.5% in controls to 15.6 ± 2.2%, 25.8 ± 4.9, and 22.9 ± 1.8% in ultraviolet-irradiated cells with no treatment, PTHrP 1–34 treatment, or PTHrP 67–86 treatment, respectively (Figure 3, *P < 0.001 control, **P < 0.01 versus irradiated with no treatment). Condensed nuclei were slightly more frequent among nonirradiated cells treated with either PTHrP peptide compared with untreated cells, but the differences were not significant. Nuclear condensation did not differ between PTHrP 1–34 treatment and PTHrP 67–86 treatment in either nonirradiated or irradiated cells.
Figure 4 demonstrates the nuclear morphology of control cells with cells exposed to ultraviolet with and without peptide treatment. Apoptotic bodies (arrowheads, representative cells) and cells with fragmented or condensed nuclei (arrows) are present among irradiated cells.
Nonadherent cells were 100% nonviable as assessed by trypan blue staining, regardless of ultraviolet exposure or treatment with PTHrP peptides (n = 3 cell preparations). Cytospin preparations of nonadherent cells were evaluated by fluorescent microscopy in comparison with cells trypsinized from normal healthy cultures. In the absence of ultraviolet exposure, a fraction of nonadherent cells had condensed or fragmented nuclei consistent with apoptosis (Figure 5B). In contrast, close to 100% of nonadherent cells from cultures exposed to ultraviolet appeared to be apoptotic (Figures 5C–5D).
Figure 6A presents time courses from representative caspase 3 assays. Fluorescence increased with time due to cleavage of Ac-DEVD-AMC, a nonfluorescent caspase 3 substrate, to fluorescent AMC. The rate of increase in fluorescence was increased in lysates of cells exposed to ultraviolet (Figure 6A) compared with control cells. Ac-DEVD-CHO, a specific inhibitor of caspase 3, dramatically decreased the cleavage rate when present in the assay mixture. On average, caspase 3 activity increased from 2.3 ± 0.4 to 5.4 ± 0.1 RFU/min in nonirradiated and irradiated untreated cells, respectively (P < 0.05, Figure 6B). PTHrP 1–34 and PTHrP 67–86 treatment caused additional increases to 8.4 ± 1.4 and 9.0 ± 1.5 RFU/min/µg protein (**P < 0.05 versus ultraviolet-untreated cells). The stimulatory effects were not significantly different between the two peptides. The peptides had no effect on caspase 3 activity in nonirradiated cells.
Activated Akt expression in type II cells after ultraviolet and PTHrP peptides. Activated Akt was present in Jurkat cell lysates but was not detected in control or ultraviolet-irradiated type II cell lysates (Figure 7). PTHrP peptide treatments had no effect.
Modulation of Ultraviolet and PTHrP Peptide Effects by Serum-Free Conditions Caspase 3 activity was increased in cells incubated with serum-free media compared with cells exposed to serum (Table 2). For example, activities in control cells without ultraviolet exposure were 2 ± 1 and 17 ± 5 RFU/min/µg protein in serum-containing and serum-free media, respectively (first column of table, first versus third lines of data). In serum-free media, PTHrP 1–34 and PTHrP 67–86 treatment augmented caspase activity by roughly 50% over levels irradiated, untreated cells, similar to the degree of augmentation in the presence of serum. The effects of the two peptides were not significantly different.
Modulation of PTHrP Peptide Effects by Substrate Caspase 3 activity increased after ultraviolet irradiation in cells grown on fibronectin, but the response to PTHrP peptides was reduced compared with cells grown on plastic substrate (Table 2, fifth line of data). PTHrP 1–34 pretreatment increased caspase 3 activity 28%, compared with 56% in cells on plastic. PTHrP 67–86 did not increase caspase 3 activity significantly compared with irradiated untreated cells on a fibronectin substrate.
Modulation of Effects of Ultraviolet and PTHrP Peptides by Type II Cell Confluence
Effect of Caspase 3 Inhibitor on Apoptosis after Ultraviolet and PTHrP Peptides
PTHrP Signaling in Type II Cells PTHrP 1–34 activates a G protein–coupled receptor and can signal through increases in cAMP or inositol phosphates (12). Because PTHP 1–34 and PTHrP 67–86 had similar effects on apoptosis, we investigated whether they augmented cAMP or inositol phosphate levels. A 10-min treatment with forskolin stimulated cAMP levels in cultured type II cells almost 15-fold compared with the basal state (Figure 9A), but PTHrP 1–34 and PTHrP 67–86 treatments had no effect. Incubation with ATP for 10 min increased inositol phosphate levels by 75 ± 20% compared with untreated type II cells (Figure 9B, P < 0.001). PTHrP 1–34 and PTHrP 67–86 augmented inositol phosphate levels by 35 ± 9 and 45 ± 17%, respectively (P < 0.05).
The initial experiments in this study investigated whether ultraviolet radiation killed cultured rat type II cells and whether the mode of death was apoptosis. The decrease in the density of adherent cells and the increase in the number of nonviable nonadherent cells (Figures 1 and 2, and Table 1) indicate that irradiation does kill type II cells, but does not identify the mode of death. Increased permeability to ethidium bromide or other vital dyes is a feature that can be common to cells dying by either apoptosis or necrosis. Therefore, we also examined several markers that are more specific for apoptosis—nuclear condensation of adherent cells, the presence of apoptotic bodies, and caspase 3 activity. The changes in these measurements (Figures 3 and 4) indicate that apoptosis is a major mode of type II cell death after ultraviolet irradiation. Our value for the extent of type II cell apoptosis with ultraviolet exposure, 16% of cells after 24 h, is comparable to results of other investigators. Edwards and colleagues observed that 20% of type II cells were apoptotic 24 h after exposure to 0.5 J/cm2 ultraviolet-C (18), whereas White and coworkers found 85% apoptosis after irradiation with a higher dose, 47 J/cm2, of ultraviolet-A (19). PTHrP sensitizes cells to apoptotic stimuli or causes apoptosis in several cell types. Activation of the type I PTH/PTHrP receptor induces apoptosis in human embryonic kidney (HEK) 293 cells (6). PTH 1–34 induces apoptosis and PTHrP 1–34 augments the reduction in cell viability induced by dexamethasone in postconfluent mesenchymal cells (5). Also, PTHrP gene transfer increases apoptosis after serum withdrawal in IEC-6 cells, a nonmalignant rat intestinal epithelial cell line (25). Our main goal was to investigate the effects of PTHrP 1–34 and PTHrP 67–86 on type II cell apoptosis after ultraviolet exposure. We found that the two peptides were proapoptotic. Treatment with either peptide augmented the reduction in type II cell viability after ultraviolet irradiation, approximately doubled the increase in nuclear condensation, and increased caspase 3 activity after ultraviolet exposure. PTHrP 1–34 and PTHrP 67–86 did not induce apoptosis by themselves over a 24-h incubation period (Figures 3, 4, and 6). PTHrP 1–34 increased membrane permeability in nonirradiated cells (Table 1), but without progression to overt apoptosis. The proapoptotic effects on type II cells in our study were elicited by treatment with exogenous peptide and thus were presumably mediated through a paracrine route. Similarly, the proapoptotic stimuli in HEK 293 cells and the mesenchymal cells were paracrine effects, mediated through the interaction of PTHrP 1–34 with its receptor. In contrast, sensitization to apoptosis in the IEC-6 cells was mediated through an intracrine effect of PTHrP at the cell nucleus. The effect was abrogated by mutations in the nuclear localization sequence of PTHrP (25). We have not investigated intracrine effects of PTHrP in type II cells.
Caspase 3 activity is a marker for apoptosis because it is a downstream executioner caspase that catalyzes the cleavage of many key enzymes and structural proteins during apoptosis, such as poly(ADP-ribose) polymerase (PARP), protein kinase C The proapoptotic effects of PTHrP might be mediated by decreasing levels of activated Akt, as PTHrP 1–34 does in postconfluent mesenchymal cells (5). Akt (also known as protein kinase B), a serine/threonine protein kinase downstream of phosphatidylinositide 3'-OH kinase (PI3 kinase), is important for survival of many cell types, including type II cells (27). We examined activated Akt levels by immunoblotting in control cells and irradiated cells with and without PTHrP 1–34 and PTHrP 67–86 treatment. Phosphorylated Akt was not detected under any of the conditions in type II cells, but was present in Jurkat cell positive controls. The absence of positive results in type II cells was not due to specificity problems, because the same antibody has been used to demonstrate increases in activated Akt in type II cells after treatment with surfactant apoprotein-A (27). We conclude that decreases in activated Akt are not of prime importance in the proapoptotic effects of PTHrP peptides after ultraviolet irradiation. We also investigated the effects of PTHrP peptides in cells grown in serum-free conditions. PTHrP might contribute to cellular demise by interfering with the action of growth factors needed for survival. If this were the case, the effects of PTHrP would be reduced in serum-free media because of the absence of these factors. However, PTHrP peptides were proapoptotic whether or not the growth factor milieu provided by serum was present, indicating that the mechanism did not depend on interactions with other factors. Apoptosis can be regulated by cell–substrate and cell–cell interactions (28). Cell–substrate effects may be mediated by integrins, heterodimeric receptors that recognize various extracellular matrix proteins (28). Ligation of integrins can activate intracellular pathways including mitogen-activated kinase cascades or focal adhesion kinase. Our study indicates that extracellular interactions can modulate the response to PTHrP in type II cells. Culture on fibronectin or growth to confluence both ameliorated the proapoptotic effects of PTHrP in ultraviolet injury. Matrix proteins have been shown to protect lung epithelial cells from injurious stimuli. Several substrates, including fibronectin, reduce alveolar type II cells DNA damage in hyperoxia and facilitate survival of bronchial epithelial cells (29, 30). Confluence protects a variety of cell types from apoptosis (31, 32), consistent with our observation of reduced proapoptotic PTHrP effect in confluent type II cells. Our results are in contrast to findings in mesenchymal cells, where confluence transforms PTHrP from an antiapoptotic to a proapoptotic factor (5). The loss of PTHrP responsiveness in type II cells could represent changes concurrent with the time-dependent alteration in phenotype that proceeds in time in primary cultures of alveolar epithelial cells. Alternatively, the effect might result from signaling pathways following cell–cell interaction. In vestibular epithelia, cell–cell contact mediates a protective effect on cell survival through interaction of E cadherin, a transmembrane cell–cell adhesion molecule, with ß-catenin (32). We included PTHrP 67–86 in our investigation because we have previously demonstrated that this peptide and PTHrP 1–34 have similar actions in stimulating phosphatidylcholine secretion in cultured type II cells and increasing bronchoalveolar lavage phosphatidylcholine levels in rats with silica lung injury (14). Thus, we hypothesized that PTHrP 67–86 might regulate type II cell apoptosis if PTHrP 1–34 did. A receptor for PTHrP 67–86 has not been identified, but biological effects are well established for this molecule. For example, PTHrP 67–86 stimulates increases in intracellular calcium and inositol triphosphates in SqCC/Yq oral buccal mucosal carcinoma cells (33). The same peptide augments or inhibits growth in various clones of 8701-BC breast carcinoma cells and stimulates ovine placental calcium transport (9, 34). Because PTHrP 67–86 stimulates inositol phosphates in SqCC/Yq cells and type II cells, one might predict that a receptor, if found, would be a G protein-coupled receptor, as is the PTHrP 1–34 receptor. We have not yet established the signal transduction pathways through which PTHrP 1–34 and PTHrP 67–86 sensitize type II cells to apoptosis, but the results of our secondary messenger production experiments suggest that the proapoptotic effects could be mediated through a phospholipase C pathway. This is the mechanism for the apoptotic effects of PTHrP 1–34 in HEK 293 cells (6). In contrast, proapoptotic effects of PTHrP 1–34 in postconfluent mesenchymal cells depend on cAMP activation and are associated with inhibition of Akt phosphorylation (5). The PTH/PTHrP receptor for which PTHrP 1–34 is a ligand can be coupled to multiple G-proteins, including Gs, Gi, and Gq, and signal transduction may vary in a cell-dependent fashion (12). Our results indicate coupling to Gq, but not necessarily to Gs. The physiologic significance of the proapoptotic effects of PTHrP remains to be determined. The effects of PTHrP on type II cell differentiation and cell death suggest that PTHrP may contribute to the natural progression of type II cells through the life cycle, i.e., maturation followed by senescence. Lung PTHrP expression changes in a regulated fashion after hyperoxic and silica-induced lung injury, probably due to changes in production by type II cells (35–37). Levels fall in the acute period after injury during the period of intense type II cell proliferation, and then rise to baseline or higher levels. The early reduction in PTHrP levels could be important for increasing the effect of the proliferative response, whereas the return to baseline could contribute to homeostatic mechanisms involved in eliminating excess pneumocytes. Further studies are indicated. In summary, we have demonstrated that ultraviolet irradiation induces a decrease in type II cell viability predominantly through apoptosis and that pretreatment with PTHrP 1–34 or PTHrP 67–86 augments the apoptotic effects of the radiation. The effects do not depend on contribution or interaction with growth factors in serum in the media,. Effects are ameliorated in confluent cells or by culture on fibronectin, suggesting interactions with signaling pathways that may be activated by cell–substrate or cell–cell interactions. In addition to effects on apoptosis, PTHrP 1–34 and PTHrP 67–86 also have similar effects on type II cell surfactant synthesis, suggesting that the two portions of the molecule act through similar mechanisms. PTHrP 1–34 has proapoptotic effects in other cell types, mediated through cAMP or phospholipase C/Cai++ pathways. The mechanisms for the effects of the amino-terminal and mid-molecule PTHrP peptides and the physiologic significance remain to be elucidated.
This work was supported by VA Merit Review grants (R.H.H. and L.J.D.), NIH grants ES09227 (R.H.H.) and DK60588 (L.J.D.), and the California Tobacco-Related Disease Research Program Grant Number 10RT-0161. Received in original form December 20, 2002 Received in final form June 3, 2003
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Abstract
Introduction
10%. The two PTHrP peptides augmented the increase in condensed nuclei 4- to 5-fold compared with nonirradiated cells, another 8–10% (*P < 0.001 versus nonirradiated controls, **P < 0.01 versus nonirradiated and irradiated control cells). Condensed or fragmented nuclei were slightly more frequent in nonirradiated cells treated with PTHrP peptides, but the changes compared with untreated cells were not statistically significant. The effects of the two PTHrP peptides were not statistically different. The numbers under the columns indicate the number of cell isolations from which the data are derived.
, and lamin A (
