Induction of Integral Membrane PAM Expression in AtT-20 Cells Alters the Storage and Trafficking of POMC and PC1

An AtT-20 cell line, engineered for inducible PAM (iPAM) expression using the rTet has been described previously (Gossen et al., 1993, 1994; Schiller et al., 1997). AtT-20/D-16v cells (Milgram et al., 1992) were initially transfected with pUHD17-2.neo, selected with G418 (0.5 mg/ml; Sigma) and screened by slot blot and Northern blot for RNA using a rtTA cDNA probe. Clonal pUHD cell lines were obtained by subcloning and screened by in situ hybridization (Schiller et al., 1997). The pUHD.PAM vector was cotransfected into AtT-20/pUHD cells with pSCEP to confer hygromycin resistance (Paquet et al., 1996). Cells containing all three plasmids were selected by growth in medium containing G418 (0.1 mg/ml) and hygromycin (200 U/ml; Sigma) and were subcloned as required. The resulting iPAM cell lines were evaluated by immunostaining with PAM antisera. Cell lines were maintained in DME/F12 medium containing sera from donor herds (10% defined equine serum; Hyclone Laboratories) in an atmosphere of 95% air, 5% CO₂. Nontransfected cells and cell lines stably expressing pUHD, PAM-1, or PHM4 were maintained as previously described (Milgram et al., 1992). A new stable PAM-1 line was also generated for this study using a different vector (pCI.neo) but with identical properties to the original cell line.

iPAM cells were treated with the antibiotic doxycycline (Dox) to induce PAM expression. The health and viability of the cells were normal up to 4 μg/ml Dox, but treatment with 8 μg/ml Dox was associated with decreased PAM protein and enzymatic activity and rounding up of the cells. To determine the optimal treatment time for Dox induction of PAM expression, iPAM cells were treated with a maximally effective dose of Dox (4 μg/ml) for varying time periods. Using antisera to PHM and PAL, increased levels of PAM protein were first detectable after 8 h of Dox treatment. Peak levels of PAM protein were observed after 48 h of Dox treatment and longer treatment of iPAM cells (72 or 96 h) did not alter the PAM expression levels in these cells (data not shown). In addition, iPAM cells treated for up to 14 d with optimal doses of Dox showed no apparent detrimental effects (data not shown). Removal of Dox from iPAM cells treated with 4 μg/ml Dox for 48 h slowly restored PAM expression to basal levels (data not shown); 24 h after removal of Dox there was no decline in PAM expression. Dox had no noticeable effect on nontransfected cells under the same conditions used for induction of iPAM cells (data not shown).

Polyclonal rabbit antisera raised to different regions of PAM were used: Ab1764 (rPAM-1[37-382]) detected PHM; Ab471 (rPAM-1[464-864]) detected PAL (Milgram et al., 1992). mAb 6E6 (rPAM-1[898-976]) detected the cytoplasmic domain of rPAM-1 and mAb 18E5 detected the PHM domain (Yun et al., 1993; Milgram et al., 1997). Other rabbit polyclonal antisera used detected the NH₂-terminal region of ACTH (JH93; Zhou et al., 1993), PC1[359-373] (JH888; Zhou and Mains, 1994), and TGN38[155-249] (JH1479; Milgram et al., 1997). Rabbit antisera were raised at Covance.

Total RNA was isolated using RNA Stat-60 reagent (TelTest B) and fractionated using denaturing formaldehyde gels. PAM mRNA was visualized using a PHM probe (1.3-kb PstI-BamHI fragment of rPAM-1, nt 356-1682; Thiele and Eipper, 1990). RNA loading was normalized using a probe for the ribosomal protein S26 (Vincent et al., 1993).

Cell extracts or conditioned media were fractionated on either 10 or 12% polyacrylamide, 0.25% N,N′-methylene-bis-acrylamide/SDS slab gels, transferred to Immobilon-P membranes (Millipore) and visualized using the enhanced chemiluminescence kit (Amersham Lifescience) (Husten and Eipper, 1991; Milgram et al., 1992). Densitometric analysis of the immunoblots was performed using NIH Image analysis software (National Institute of Mental Health) (Milgram et al., 1993).

Cells were scraped into ice-cold 20 mM Na(N-tris[hydroxymethyl]methyl-2-aminoethanesulfonic acid) (NaTES)/10 mM mannitol, pH 7.4, 1% Triton X-100 (TMT), containing protease inhibitors (Milgram et al., 1992). After three cycles of freezing and thawing, the supernatant was retained and the pellet was discarded. Protein concentrations were determined using the bicinchoninic acid protein reagent kit (Pierce Chemical Co.). PHM and PAL assays were performed on cell extracts as previously described using acetyl-Tyr-Val-Gly and acetyl-Tyr-Val-α-hydroxyglycine as substrates, respectively (Kolhekar et al., 1997).

Cells were plated on 12-mm culture dishes coated with 0.1 mg/ml poly- l-lysine (Sigma) and grown for 48 to 72 h before biosynthetic labeling. Cells were incubated for 10 min in complete serum-free medium (CSFM) lacking methionine (selectamine kit; GIBCO BRL), then labeled for 30 min with 250 μCi [³⁵S]methionine (∼1 μM [Met]; Amersham) at 37°C in air atmosphere. Cells were chased in CSFM containing 140 μM methionine, 2 mg/ml BSA, 0.1 mg/ml lima bean trypsin inhibitor, pH 7.4, for different time periods. After the chase periods, the spent medium was collected, centrifuged at 2,000 g to pellet nonadherent cells, and the supernatant was transferred to a new tube containing a cocktail of protease inhibitors (Milgram et al., 1992). All samples were kept at −80°C until further analysis. Cell extracts (5 × 10⁶ dpm, trichloroacetic acid precipitable) and equivalent amounts of spent media were subjected to immunoprecipitation using a rabbit polyclonal antiserum to PAM. Immune complexes were collected with protein A beads and analyzed by SDS-PAGE followed by fluorography (Zhou et al., 1993).

Duplicate wells of cells (nontransfected and stably transfected PAM-1, iPAM−, iPAM+, pUHD−, and pUHD+) were plated on poly-l-lysine– treated wells for measurements of ACTH secretion. Wells were initially rinsed for three 30-min periods in basal release medium (DME/F12/Air with 2 mg/ml fatty acid–free BSA, 0.1 mg/ml lima bean trypsin inhibitor, 1 μg/ml insulin, 0.1 μg/ml transferrin) (Mains and Eipper, 1981; Milgram et al., 1993). The experiment was begun with two successive 30-min basal collections taken using control medium and concluded with one 30-min period of stimulated secretion (1 mM BaCl₂ in basal release medium). The collected medium was centrifuged to remove nonadherent cells and protease inhibitors were added and the medium frozen until assay. Cells were extracted using 5 N acetic acid with protease inhibitors, lyophilized, and then dissolved in radioimmunoassay buffer with protease inhibitors. ACTH radioimmunoassays were performed using Ab Kathy (1:15,000) and [¹²⁵I]ACTH(1-39) (Amersham); this antiserum only recognizes POMC products in which the COOH-terminal end of ACTH(1-39) is exposed (Schnabel et al., 1989).

To detect filamentous actin, cells were fixed in prewarmed 4% paraformaldehyde for 20 min, washed once with PBS (50 mM sodium phosphate, 150 mM sodium chloride, pH 7.4), permeabilized for 20 min (0.075% Triton X-100, 2 mg/ml BSA in PBS), washed once, and incubated in block for 30 min (2 mg/ml BSA in PBS). Cells were incubated with a PAM mAb (6E6) for 120 min, then washed three times with PBS. PAM staining was detected with a donkey anti–mouse Cy3 and filamentous actin visualized with a 1.2 μg/ml FITC-phalloidin. The cells were exposed to this second set of antibodies for 60 min (in the dark), washed three times in PBS, and then mounted with prewarmed DABCO/permount and coverslipped.

Cells were plated on poly-l-lysine–coated glass chamber slides (Lab-Tek) 48–96 h before fixation. Cells were double immunostained for PAM and ACTH, PAM and TGN38, or for PAM and PC1 as described (Milgram et al., 1997). Cells were viewed under epifluorescence optics with an upright microscope (Axioskop; Zeiss, Inc.) using FITC (BP 485/20, barrier filter 520-560) and rhodamine (BP 546/12, LP 590) filters with an HBO 100-W mercury lamp and a ×40 or ×63 objective lens. Images were photographed using a Micromax digital camera with Winview 3.2 software (Princeton Instruments). One set of images was visualized on a confocal laser microscope (Noran OZ; Noran) at an excitation wavelength of 488 nm (FITC-nar, barrier filter 525 ± 50) and 568 nm (Rho-nar, barrier filter 605 ± 55) from a Krypton-Argon multi-line laser. Z-axis images were processed using Intervision 1.6 software (Noran). A second set of images was visualized on a confocal laser microscope (MRC 600; Bio-Rad Laboratories) at an excitation wavelength of 488 nm (FITC-nar, barrier filter 510-515) and 568 nm (Rho, barrier filter 585) from a Krypton-Argon laser. Z-axis images were processed using Comos 6.05 software. Photographs of nontransfected and transfected cells were taken under identical conditions and printed in an identical fashion. A set of control experiments (omission of primary antibody or omission of secondary antibody) was performed showing that there was no bleed through from one fluorophore into the other filter when appropriate dilutions of antisera were used for double immunostaining.

Induction of PAM-1 mRNA was examined by Northern blot (Fig. 2). iPAM cells were treated with increasing concentrations of Dox for 48 h. In the exposure shown, there is no detectable PAM mRNA in the absence of Dox and the intensity of the signal increases with higher doses of Dox, reaching maximal expression levels at 1–4 μg/ml Dox. With longer exposure times, the level of PAM mRNA in the noninduced iPAM cells was found to be similar to the level of endogenous PAM mRNA in the nontransfected cells (data not shown). There was a 60-fold induction of PAM mRNA expression in the maximally induced iPAM cells compared with the noninduced cells. No decrease in PAM-1 mRNA level was observed with the highest dose of Dox (8 μg/ml). The level of PAM-1 mRNA in the maximally induced cells was similar to the level in stably transfected PAM-1 cells (data not shown).

Since one of our goals was to examine cells expressing different levels of PAM-1, iPAM cells were treated for 48 h with increasing concentrations of Dox and compared with control cells using enzyme assays (Fig. 3 A) and Western blots (Fig. 3, B and C). Controls included nontransfected AtT-20 cells as well as AtT-20 cells expressing only the reverse tetracycline repressor (pUHD−, untreated; pUHD+, treated with 4 μg/ml Dox for 48 h) or expressing PAM-1 or PHM4. As with PAM mRNA (Fig. 2), Dox exerted a dose-dependent effect on PHM activity (Fig. 3 A). Peak levels of PHM specific activity were observed after treatment with 1–4 μg/ml of Dox. PHM specific activity in the noninduced iPAM cells was similar to nontransfected cells, untreated, and Dox-treated pUHD cells (Fig. 3 A). The PHM specific activity of maximally induced iPAM cells was 20–30-fold higher than noninduced iPAM cells and comparable to that of the stably transfected PAM-1 and PHM4 cells (Fig. 3 A). The sensitivity of iPAM cells to induction with low amounts of Dox can be seen by the two- to fivefold increase in PHM specific activity when noninduced cells are compared with cells treated with 0.0625 μg/ml Dox.

Increasing concentrations of Dox produced a dose-dependent increase in PAM protein expression in the iPAM cells (Fig. 3 B). Intact PAM-1 (120 kD) and soluble PHM (46 kD) were apparent in the Dox-treated iPAM cells as in the stably transfected PAM-1 cells (Fig. 3 B). Since 46-kD PHM can be stored or secreted, spent media were also examined (Fig. 3 C). Cells were incubated in CSFM for 6 h; equal amounts of cell protein (Fig. 3 B) and a 10-fold higher volume of spent medium (representing secretion) (Fig. 3 C) were visualized with an antiserum to PHM. Due to the limited sensitivity of our antisera, endogenous PAM expression in iPAM cell extracts before induction (0 Dox) or in nontransfected cells, pUHD−, and pUHD+ cells is not detectable (Fig. 3 B). The addition of Dox (4 μg/ml for 48 h) to the culture medium of nontransfected and PAM-1 cells had no effect on PAM protein expression (data not shown).

Maximum PAM protein levels in the iPAM cell line (1–4 μg/ml Dox) were similar to levels in the stably transfected PAM-1 cell line and at least 35–40-fold higher than in the noninduced iPAM cells. Western blots of cell extracts and media were digitized (Fig. 3, D and E). In the iPAM cells, the proportion of 46-kD PHM to the total amount of PAM protein in the cell extract decreased 37% with increasing PAM protein expression (Fig. 3 D). The decreased fraction of 46-kD PHM with increased total cellular PAM expression could reflect decreased cleavage of PAM-1 or increased secretion of 46-kD PHM.

Secretion of PHM from iPAM cells was detectable with Dox doses of 0.0625 μg/ml or higher (Fig. 3 E). At the lowest doses of Dox, more 46-kD PHM was in the cells (filled bars) than in the medium (open bars) after the 6-h basal collection. AtT-20 cells producing soluble PHM (PHM4) store this protein much more efficiently than AtT-20 cells that produce soluble PHM from membrane PAM. As expression of PAM-1 increases, the recovery of 46-kD PHM decreased (Fig. 3 D) and the fraction of the 46-kD PHM recovered in the medium increased (Fig. 3 E). As a result, the amount of secreted 46-kD PHM was similar at the higher levels of induction (>2 μg/ml Dox). The decrease in steady-state levels of 46-kD PHM with increasing levels of PAM-1 expression demonstrates that there are saturable steps in PAM-1 metabolism. In addition, the ability of the cells to store the 46-kD PHM that is produced declines as expression of PAM-1 increases.

As with the PAM mRNA, PHM protein expression, and PHM enzyme activity, a dose-dependent effect of Dox on PAL enzyme activity was observed (Fig. 4 A). Increased expression of PAL activity was apparent with the lowest doses of Dox tested. Peak levels of PAL specific activity were observed after treatment with 2–4 μg/ml Dox; treatment with 8 μg/ml Dox resulted in decreased PAL activity. PAL specific activity in the noninduced iPAM cells was similar to nontransfected, untreated, and Dox-treated pUHD cells (Fig. 4 A). The PAL specific activity of maximally induced iPAM cells was comparable to that of the stably transfected PAM-1 cells and two- to threefold higher than PHM specific activity, reflecting the higher enzymatic turnover rate of PAL (Eipper et al., 1993).

Processing of PAM-1 in the iPAM cells yields a 70-kD PALm product, as in stably transfected PAM-1 cells (Fig. 4 B). A dose-dependent increase in PAM protein expression was seen in the iPAM cells using the PAL antiserum, with levels approaching those in PAM-1 cells; very little soluble PAL is produced from PAM-1 in AtT-20 cells, so Western blots of PAL in spent media are not shown. Western blots were digitized so levels of expression could be compared (Fig. 4 C). The fraction of the total PAM protein in the cell extracts accounted for by 70-kD PALm increased twofold with increasing PAM expression levels (Fig. 4 D). PAM-1 metabolism is clearly altered as a function of expression level.

The alterations in steady-state levels of PHM and PAL with increasing levels of PAM expression could reflect differences in the rate of biosynthesis of PAM-1, cleavage of PAM-1, degradation, and/or secretion of PHM and PAL. To distinguish these possibilities, identical wells of iPAM cells treated with a low dose of Dox (0.25 μg/ml) or a high dose of Dox (2.0 μg/ml) were incubated in medium containing [³⁵S]Met for 30 min and either harvested immediately (pulse) or chased in medium containing unlabeled Met (chase). PAM proteins immunoprecipitated from cell extracts and media were fractionated by SDS-PAGE (Fig. 5). Consistent with the steady-state levels of PHM and PAL protein and enzyme activity (Figs. 3 and 4), the rate of PAM-1 biosynthesis was 2.8 ± 0.4-fold higher (P < 0.005; n = 4) in cells induced with the high dose of Dox than in cells induced with the low dose of Dox.

Consistent with Western blot analysis (Fig. 3), relative secretion of 46-kD PHM was enhanced when levels of PAM-1 expression were higher. With the low dose of Dox, 65% of the 46-kD PHM present after a 4-h chase was recovered in the medium. In contrast, with the high dose of Dox, 81% of the 46-kD PHM was recovered in the medium. No detectable degradation of PAM protein occurred within 4 h at either level of PAM expression. The higher levels of 70-kD PALm observed in PAM-1 cells induced with high levels of Dox may reflect a small change in the turnover rate of this membrane protein.

To evaluate the localization of PAM at different levels of expression, a dose–response curve for Dox induction of PAM-1 in iPAM cells was examined by immunofluorescence (Fig. 6 A). Cells were fixed and visualized with a mAb directed against the PAM-1 COOH terminus. Less than 1% of the cells stained for PAM-1 when the untreated iPAM cells were examined (Fig. 6 A, 0 Dox); as for Western blots, our antisera cannot detect the endogenous levels of PAM in AtT-20 cells. In iPAM cells treated for 48 h at a low Dox dose (0.25 μg/ml), PAM-1 signal was seen in ∼50% of the cells (Fig. 6 A, 0.25 Dox). In maximally induced iPAM cells (4 μg/ml Dox), PAM staining was much more intense and >95% of the cells displayed fluorescence (Fig. 6 A, 4 Dox).

Maximally induced iPAM cells were coimmunostained with antisera for a TGN marker protein (TGN38) and PAM-1 (Fig. 6 B). In the panels shown, two cells show intense PAM-1 staining (green) in the perinuclear region with lighter staining in vesicular structures at the tips of processes (arrows). When the PAM-1 and TGN38 images were superimposed, the TGN region was predominantly yellow, indicating that much of the PAM-1 is localized to the TGN region (Fig. 6 B); TGN staining was not apparent at the tips of the processes (arrows). In a third cell that stained only faintly for PAM-1, the TGN staining pattern was similar to the staining pattern seen in cells expressing more PAM-1 (Fig. 6 B).

Having established that the iPAM cells can be used effectively to control the expression of PAM in a given cell population, the effects of increased PAM expression on ACTH localization were evaluated by immunofluorescence (Figs. 6 C and 7). Fig. 7 shows epifluorescence images of populations of nontransfected and PAM-1–expressing cells whereas Fig. 6 C shows confocal images of two cells among a population of iPAM cells treated with Dox; as occasionally happens, only one of the pair of cells is expressing high levels of PAM (green). In the population of nontransfected (NT) cells (Fig. 7, left), vesicular, punctate staining for ACTH is observed throughout the cytosol with increased staining in the cellular processes and a slight increase in ACTH staining in the TGN region. In the single iPAM cell not expressing high levels of PAM (Fig. 6 C, left cell), the ACTH (red) staining pattern is similar to that of the nontransfected cells. pUHD cells treated with Dox also displayed ACTH staining patterns similar to those seen for nontransfected cells (data not shown). Interestingly, in the confocal image of the iPAM cell expressing PAM-1 (Fig. 6 C, right), the ACTH distribution is dramatically altered, with marked localization of ACTH to the TGN region of the cell and less intense staining of processes. Similar patterns of ACTH staining are observed in the population of AtT-20 cells expressing PAM-1 (Fig. 7, middle). The PAM and ACTH staining patterns of the Dox-treated iPAM cells (Fig. 6 C, right cell) are almost identical to those of the stably transfected PAM-1 cells (Fig. 7, PAM-1).

Confocal microscopy allowed more detailed comparison of the localization of ACTH and PAM in the iPAM+ cells. PAM staining in this cell (Fig. 6 C) is predominantly localized to the TGN region with some staining for PAM at the tips of the cellular processes. When the ACTH and PAM signals are superimposed, a large portion of the TGN region is yellow, consistent with colocalization of ACTH with PAM-1. Compared with the noninduced iPAM cell (Fig. 6 C, left cell), the intensity of ACTH staining is decreased in the cell with increased PAM expression (Fig. 6 C, right cell); a similar difference in staining intensity is seen when comparing ACTH staining in nontransfected and PAM-1 cells (Fig. 7). The different ACTH localization observed in iPAM cells expressing PAM-1 cannot be caused simply by the introduction of proteins into the secretory pathway using the rTet system, since previous results using AtT-20 cells with Dox-inducible expression of regulated endocrine–specific protein of 18 kD showed no effect on POMC biosynthesis or localization (Schiller et al., 1997).

To determine which region of PAM was responsible for altering ACTH localization, nontransfected and stably transfected AtT-20 cell lines expressing either intact PAM-1 or PHM4 (Fig. 7) were compared. In cells expressing PHM4 (lacking the PAL and transmembrane domains of PAM-1), PHM staining differs from that of PAM-1 and iPAM+ cells in that it is primarily seen in the cellular processes with fainter staining in the TGN region (Fig. 7, PHM4). Moreover, the ACTH staining pattern in the PHM4 cells closely resembles that of the nontransfected and noninduced iPAM cells (Figs. 7 and 6 C, left cell, respectively); ACTH staining in the PHM4 cells is primarily seen in the cellular processes with less staining of the TGN region.

Observing that the ACTH immunostaining patterns were markedly different in nontransfected cells and cells expressing PAM-1 (Figs. 6 C and 7), their basal and stimulated secretions of ACTH were examined. Secretion of ACTH was quantified using an immunoassay specific for the COOH terminus of ACTH(1-39); secretion was examined during two sequential basal collections and cells were then stimulated with 1 mM BaCl₂ (Fig. 8 A). BaCl₂ mimics Ca²⁺ and several secretagogues and supports a steady high rate of secretion (Mains and Eipper, 1984). As expected, in nontransfected AtT-20 cells, the addition of secretagogue to the medium stimulated ACTH secretion three- to fourfold above basal levels. A similar fold stimulation of ACTH secretion was observed in control pUHD cells whether or not they were treated with Dox (pUHD−, pUHD+) and in noninduced iPAM cells; none of these cells express exogenous PAM-1. In contrast, in iPAM cells treated with Dox to induce PAM-1 expression, addition of secretagogue to the medium did not stimulate ACTH secretion above basal levels (Fig. 8 A). The effects seen with the iPAM cells cannot be attributable to the Dox treatment alone since the pUHD+ cells responded to secretagogue as well as nontransfected AtT-20 cells or untreated pUHD− cells. As for Dox-treated iPAM cells, secretion of ACTH from stably transfected PAM-1 cells was not stimulated by secretagogue (Fig. 8 A). In contrast, AtT-20 cells expressing a soluble PAM protein (PHM4), exhibited a threefold increase in ACTH secretion in response to secretagogue. Treatment of the iPAM cells with a low, but effective dose of Dox (0.0625 μg/ml) resulted in a partial inhibition of the ability of secretagogue to stimulate ACTH secretion (data not shown). Thus, overexpression of PAM-1 is causally related to the inability of Dox-treated iPAM cells and PAM-1 cells to respond to secretagogue treatment.

In the course of analyzing ACTH secretion by the various AtT-20 cell lines, the utility of having an inducible cell line was clearly apparent (Fig. 8 B). The pUHD cell line selected contained roughly half as much ACTH per milligram of cell protein as the parental, nontransfected AtT-20 cell line. The iPAM cell line subsequently selected contained roughly half as much ACTH per milligram of cell protein as the parental pUHD cell line. In no case did Dox treatment affect the ACTH content of the cells. The PAM-1 and PHM4 AtT-20 lines generated several years ago contained approximately as much ACTH per milligram of cell protein as the newly generated iPAM cell line.

To evaluate regulated secretion of another lumenal protein, the level of PHM activity was determined in the same set of basal and secretagogue stimulated media samples (Fig. 8 C). In the nontransfected AtT-20 cells, the level of PHM activity in the media was sixfold higher after secretagogue stimulation. A similar fold stimulation of secreted PHM activity was observed after secretagogue stimulation of control pUHD cells (whether or not they were treated with Dox [pUHD−, pUHD+]), noninduced iPAM cells, and PHM4 cells. As seen for the ACTH immunoreactivity (Fig. 8 B), cells with increased exogenous PAM-1 expression (iPAM+ and PAM-1 cells) failed to show a robust stimulation of secretion of PHM activity after secretagogue stimulation (only double the basal levels).

To ask whether expression of membrane PAM might affect other LDCV proteins in addition to ACTH, PC1 was investigated. The major products of proPC1 cleavage are mature 82-kD PC1, produced rapidly in the endoplasmic reticulum, and 66-kD PC1ΔC, produced only in LDCVs (Fig. 9 A; Zhou and Mains, 1994). A dose–response curve for Dox induction of PAM-1 expression was generated as described in Figs. 2–4 and cell extracts were subjected to Western blot analysis with antibodies to PC1 (Fig. 9 B). In nontransfected, PHM4, and pUHD− or pUHD+ cells, PC1ΔC was the major form of PC1 in cell extracts. In contrast, in PAM-1 cells, mature 82-kD PC1 predominated in cell extracts. Increasing PAM-1 expression in iPAM cells caused a dose-dependent decrease in the production of PC1ΔC from mature PC1; whereas 65% of the total PC1 was PC1ΔC in noninduced cells, only 30% was PC1ΔC in the maximally induced iPAM cells (Fig. 9 C). PC1ΔC accounted for even less of the total PC1 in stably transfected PAM-1 cells.

PC1 localization was then examined in various PAM cell lines by immunofluorescence (Fig. 9 D). PC1 staining in nontransfected and in noninduced iPAM cells was vesicular and predominantly localized to the tips of the cellular processes (Fig. 9 D, NT and iPAM−). In iPAM cells treated with Dox and expressing high levels of PAM, PC1 was not concentrated at the tips of the processes and there was an overall decrease of PC1 staining intensity. Lack of staining at the tips of processes and decreased PC1 staining intensity were also observed in stably transfected PAM-1 cells. In contrast to the iPAM+ and PAM-1 cells, the PHM4 cells showed prominent PC1 staining at the tips of the cellular processes; the PC1 staining appeared to be colocalized with the PHM staining (Fig. 9 D, PHM4). Thus the PC1 staining pattern of the PHM4 cells more closely resembles that of the nontransfected and iPAM− cells (Fig. 9 D) than the other cells tested.

Cytoskeletal organization is a powerful factor in regulated exocytosis (Muallem et al., 1995; Carbajal and Vitale, 1997). Since PAM-1 is known to interact with proteins that can indirectly affect cytoskeletal organization (Alam et al., 1997), the distribution of filamentous actin in noninduced and induced iPAM cells was examined using the mushroom toxin, phalloidin (Fig. 10). In noninduced iPAM cells expressing only their endogenous PAM protein, PAM staining was not visible (Fig. 10, iPAM−, red). Filamentous actin was found in clusters broadly distributed throughout the cell and was collected in several subplasma membrane foci (Fig. 10, iPAM−, green); although the edges of the noninduced cells were easily visible, filamentous actin was not enriched in the subplasma membrane region. As expected, after induction with Dox, PAM-1 was largely localized to the TGN region of the cell (Fig. 10, iPAM+, red). A dramatic change in the distribution of filamentous actin accompanied expression of PAM-1 (Fig. 10, iPAM+, green); filamentous actin was concentrated at the plasma membrane with numerous foci of more intense staining. The broadly distributed diffuse patches of filamentous actin were absent from the iPAM+ cells. Patches of filamentous actin were also concentrated in the TGN region of the iPAM+ cells.

PAM is an essential bifunctional enzyme that catalyzes the bioactivation of many peptides (Eipper et al., 1993). The anterior pituitary and atrial myocytes are rich sources of PAM expression (May et al., 1990; Eipper et al., 1993), whereas AtT-20 cells contain lower albeit sufficient levels of PAM to amidate the endogenous substrates. Various forms of PAM have been expressed successfully in AtT-20 cells, with stable expression of membrane PAM reaching levels found in the anterior pituitary (May et al., 1990). The use of stable cell lines is an important tool in addressing many questions relating to peptide processing and sorting, but their limitations include clonal variability and inability to regulate the level of protein expression. This problem is illustrated by the variable ACTH content of the nontransfected, PAM-1, and PHM4 cell lines (Fig. 8 B). To overcome these limitations, an inducible expression system for integral membrane PAM was developed using the tetracycline repressor system (Gossen et al., 1993, 1994; Schiller et al., 1997). By regulating the level of PAM expression in the iPAM cell line, we were able to show that the metabolism of membrane PAM is altered as expression levels increase: cleaved PHM products are less prevalent and the soluble products made are stored less well. Individual steps in PAM trafficking, such as internalization from the cell surface, remain to be evaluated.

Maximal expression of PAM in the iPAM cell line was comparable to levels in the anterior pituitary (May et al., 1990) and in cell lines stably transfected with various forms of PAM (Figs. 3 and 4). Before induction, the iPAM cells resembled nontransfected cells in their levels of PAM mRNA, enzyme activity, protein expression, and immunofluorescence. The level of PAM induction observed, when comparing nontransfected cells to maximally expressing iPAM cells, was 20–60-fold depending on the analytical method used. A 35-fold induction of chloramphenicol acetyltransferase was observed using a metallothionein-I promoter in AtT-20 cells, but the requisite levels of divalent metals had toxic effects on the cells (Dickerson et al., 1989). Taken together, these findings highlight the value of this Tet-On cell line in studying the steps in PAM metabolism by regulating its level of expression.

rPAM-1 contains several paired basic potential proteolytic cleavage sites and cleavage at the Lys-Lys⁴³⁷ site, located within the noncatalytic exon A region, results in separation of the two catalytic domains (Eipper et al., 1993); the 46-kD PHM domain is soluble whereas the 70-kD PAL domain remains membrane bound (Fig. 1; Stoffers et al., 1989). At low levels of PAM-1 expression, a higher proportion of the total PAM is 46-kD PHM and this soluble protein is largely stored in the cells. As levels of PAM-1 expression increase, decreasing amounts of the total PAM are recovered as soluble PHM, and more of the soluble PHM recovered is in the medium. The diminished cleavage of PAM-1 may reflect the fact that less 66-kD PC1ΔC is observed in cell extracts after induction of PAM-1 expression (Fig. 9). PC1 can cleave PAM-1 to soluble PHM and PALm (Marx and Mains, 1997) and PC1ΔC is more active, under the conditions in maturing LDCVs, than is 82-kD PC1 (Zhou and Lindberg, 1994). It is striking that soluble PHM produced from PHM4, a soluble precursor, is efficiently stored whereas soluble PHM produced from membrane PAM-1 is not efficiently stored (Fig. 3 C) (Milgram et al., 1994). Upon induction of PAM-1 expression in the iPAM cells, this phenomenon is replicated.

The signals mediating the trafficking of soluble, lumenal proteins and membrane proteins from TGN into immature secretory granules (ISGs) and the maturation of secretory granules are not yet understood. Our observation that induction of PAM-1 expression causes dramatic changes in the actin cytoskeleton, eliminates regulated exocytosis, causes the relocalization of soluble lumenal proteins, and limits the cleavage of lumenal proteins, provides new insight into this process. PAM-1 has lumenal and cytosolic components, and both may contribute to the observed effects. We suggest two ways in which membrane PAM could affect the regulated secretory pathway: first, the COOH-terminal domain of PAM may interact with cytosolic factors involved in regulating microtubules and the actin cytoskeleton; second, the lumenal domains of PAM may aggregate with other lumenal proteins (Colomer et al., 1996) (Fig. 11).

A working model for the effect of membrane PAM on the regulated secretory pathway in AtT-20 cells is shown in Fig. 11. The biogenesis of ISGs requires segregation of stored products from proteins leaving via constitutive-like secretion and may involve pH- and Ca²⁺-dependent selective aggregation (Kuliawat and Arvan, 1992; Kuliawat et al., 1997; Thiele et al., 1997; Kromer et al., 1998) as well as sorting receptors (Cool et al., 1997; Normant and Loh, 1998). Soluble proteins (ACTH, PC1, and soluble PHM) will aggregate as a function of the pH and [Ca²⁺] in the ISGs (Chanat et al., 1991; Colomer et al., 1996; Thiele et al., 1997). The extent of proteolytic processing may affect the ability of the proteins to aggregate (Quinn et al., 1991). In cells expressing only endogenous PAM (nontransfected, pUHD, and iPAM−) or PHM4, the prominent granular staining for ACTH and PC1 in the cellular processes (Figs. 6, 7, and 9) and the stimulation of ACTH secretion (Fig. 8 A) indicate that these soluble proteins are sorted into the regulated secretory pathway and stored in mature secretory granules (Mains and May, 1988; Schnabel et al., 1989; Milgram et al., 1994).

ISGs are situated near the TGN and clathrin-coated regions are commonly seen on them, while mature secretory granules lack clathrin-coated regions and are localized to the periphery of the cell (Tooze and Tooze, 1986; Tooze et al., 1991; Dittie et al., 1996; Klumperman et al., 1998). During the process of maturation, secretory granules become responsive to secretagogue addition; this process may involve removal of some proteins and addition of others. Retrieval of furin from ISGs involves AP-1 adaptors and clathrin (Dittie et al., 1997). Similarly, PAM-1 may be retrieved from ISGs through interactions with coat proteins. Since very little PAM-1 appears on the cell surface, whereas a significant amount of soluble PHM undergoes constitutive-like secretion, the soluble and membrane proteins leaving the ISGs must enter a compartment that allows them to part company (Fig. 11).

One of the most dramatic effects of PAM-1 expression is alteration of the actin cytoskeleton. The cytosolic domain of PAM is known to interact with Kalirin, a GDP/ GTP exchange factor for Rac1 (Alam et al., 1997). When expressed in fibroblasts, Kalirin disrupts actin stress fibers and dramatically alters cell morphology (Mains et al., 1998). Although expression of Kalirin is restricted to the nervous system in the adult rat, we postulate the existence of a Kalirin-like protein in AtT-20 cells. PAM-1, through its interaction with this Kalirin-like protein, could activate Rac1, a known effector of the actin cytoskeleton (Lamaze et al., 1997; Bellanger et al., 1998). Changes in filamentous actin can enhance or inhibit exocytosis (Muallem et al., 1995; Carbajal and Vitale, 1997; Baldini et al., 1998). In addition, the cytosolic domain of PAM interacts with P-CIP2, a serine/threonine protein kinase expressed in AtT-20 cells, that phosphorylates stathmin, a cytosolic protein known to regulate microtubule stability (Curmi et al., 1997).

Another dramatic effect of expressing PAM-1 is the complete elimination of the ability of AtT-20 cells to secrete in response to addition of secretagogue; this effect was observed in the iPAM cells and in the stably transfected PAM-1 cells (Fig. 8, A and C). Expression of soluble PHM4 did not impair the ability of AtT-20 cells to respond to secretagogue. The altered polymerization of actin may contribute to this effect in several different ways. First, the more extensive network of polymerized actin in the subplasma membrane region may serve as a physical barrier to exocytosis. Second, the altered cytoskeletal organization observed in cells expressing membrane PAM may be responsible for the failure of secretory granules to accumulate in cell processes, in a position from which exocytosis is possible. While it is clear that ISGs cannot respond to secretagogue as well as mature secretory granules, the mechanisms underlying acquisition of secretagogue responsiveness are not clear. Overexpression of PAM-1 may also impair this process.

Induction of PAM-1 expression in iPAM cells and overexpression of PAM-1 in stably transfected cells both result in the impaired endoproteolytic cleavage of POMC, PC1, and PAM itself, yielding a lumenal compartment full of partially cleaved products (POMC data not shown here; Mains et al., 1991). Maturation of the lumenal contents of the ISG is impaired. Expression of soluble PHM4 does not impair cleavage of lumenal content proteins or storage of ACTH and PC1 in vesicles that accumulate at the tips of cell processes. This effect of PAM-1 could involve both lumenal and cytosolic components. In particular, the ability of the lumenal domains of PAM to aggregate with other lumenal proteins could be important. Although the effects of overexpressing PAM-1 are widespread and diverse, they appear to be focused on the regulated secretory pathway. Localization of endogenous membrane proteins identifying the TGN (TGN38), lysosomes (LAMP1), or endosomes (mannose-6-phosphate receptor) is not dramatically altered in iPAM+ or PAM-1 cells (data not shown).

The dramatic effects of inducing PAM-1 expression in a single cell line demonstrate how well the endoproteolytic events in the lumen of the secretory granules are integrated with the maturation of ISGs and regulated exocytosis. The importance of the cytoskeleton to each of these processes is also apparent. The inducible iPAM cell line should allow us to identify the signaling pathways linking PAM-1 to the cytoskeleton, the manner in which expression of PAM-1 blocks regulated exocytosis, and the steps in the trafficking of PAM that are saturable.

We thank Drs. Manfred Bujard and Hermann Gossen for the rTet inducible vector system. We gratefully acknowledge Marie Bell and Cathy Caldwell for general laboratory assistance.

This work was supported by National Institutes of Health grants DK32948 (R.E. Mains) and DA10478 (M.R. Schiller). G. Ciccotosto was supported by a fellowship LT-647/97 from the Human Frontiers Science Program Organization.

CSFM

complete serum-free medium

Dox

doxycycline

iPAM

inducible PAM

ISG

immature secretory granule

LDCV

large dense core vesicle

NT

nontransfected

PAL

peptidyl-α-hydroxylglycine α-amidating lyase

PAM

peptidylglycine α-amidating monooxygenase

PC1

prohormone convertase 1

PHM

peptidylglycine α-hydroxylating monooxygenase

POMC

proopiomelanocortin

rTet

reverse tetracycline repressor system

TGN

trans-Golgi network