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Human Molecular Genetics, 2003, Vol. 12, No. 13 1475-1484
DOI: 10.1093/hmg/ddg160
© 2003 Oxford University Press

Protein kinase-A activity in PRKAR1A-mutant cells, and regulation of mitogen-activated protein kinases ERK1/2

Audrey Robinson-White1, Thomas R. Hundley2, Miriam Shiferaw1, Jérôme Bertherat3, Fabiano Sandrini1 and Constantine A. Stratakis1,*

1Section on Endocrinology and Genetics, Developmental Endocrinology Branch, NICHD, Bethesda, MD 20892, USA, 2LMI, NHLBI, National Institutes of Health, Bethesda, MD 20892, USA and 3Département d'Endocrinologie, Institut Cochin, INSERM U576, CNRS UMR 8104, IFR116, Université-Paris V, Paris, France

Received February 5, 2003; Revised April 17, 2003; Accepted April 28, 2003


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Carney complex (CNC) is caused by PRKAR1A-inactivating mutations. PRKAR1A encodes the regulatory subunit type I-{alpha} (RI{alpha}) of the cAMP-dependent kinase (PKA) holoenzyme; how RI{alpha} insufficiency leads to tumorigenesis remains unclear. In many cells PKA inhibits the extracellular receptor kinase (ERK1/2) cascade of the mitogen-activated protein kinase (MAPK) pathway leading to inhibition of cell proliferation. We investigated whether the PKA-mediated inhibitory effect on ERK1/2 is affected in CNC cells that carry germline PRKAR1A mutations. PKA activity both at baseline and after stimulation with cAMP was augmented in cells carrying mutations. Quantitative message analysis showed that the main PKA subunits expressed were type I (RI{alpha} and RIß) but RI{alpha} was decreased in mutant cells. Immunoblot assays of ERK1/2 phosphorylation by the cell- and pathway-specific stimulant lysophosphatidic acid (LPA) showed activation of this pathway in a time- and concentration-dependent manner that was prevented by a specific inhibitor. There was a greater rate of growth in mutant cells; forskolin and isoproterenol inhibited LPA-induced ERK1/2 phosphorylation in normal but not in mutant cells. Forskolin inhibited LPA-induced cell proliferation and metabolism in normal cells, but stimulated these parameters in mutant cells. These data were also replicated in a pituitary tumor cell line carrying the most common PRKAR1A mutation (c.578del TG), and an in vitro construct of mutant PRKAR1A that was recently shown to lead to augmented PKA-mediated phosphorylation. We conclude that PKA activity in CNC cells is increased and that its stimulation by forskolin or isoproterenol increases LPA-induced ERK1/2 phosphorylation, cell metabolism and proliferation. Reversal of PKA-mediated inhibition of this MAPK pathway in CNC cells may contribute to tumorigenesis in this condition.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Carney complex (CNC) is a multiple neoplasia syndrome characterized primarily by spotty skin pigmentation, endocrine and other tumors (1). The genes responsible for CNC have been mapped to chromosomes 2 and 17 (2). It is now known that PRKAR1A, a gene that codes for the RI{alpha} subunit of the cAMP-dependent protein kinase A (PKA) is mutated in almost half of the known patients with CNC (3). The finding of tumor-specific loss of heterozygosity (LOH) within the q22–24 region of chromosome 17 (4), and the complete or partial loss of RI{alpha} in CNC tumors suggested that RI{alpha} may function as a tumor suppressor gene in these tissues.

The PKA holoenzyme is a heterotetramer consisting of two homodimers of two regulatory (R) subunits (RI-{alpha} or -ß, and RII-{alpha} or -ß) and two catalytic (C) subunits (-C{alpha}, -Cß or -C{gamma}). PKA may be activated indirectly by stimulation of a G-protein coupled receptor, and/or the plasma membrane-located adenyl cyclase; or directly by cAMP. The binding of four cAMP molecules to two R subunits (two per each R subunit) causes the release of two free active C subunits (5), which, in turn, phosphorylate downstream targets (6).

Mitogen-activated protein kinase (MAPK), arranged in separate but interacting pathways, mediate a diverse number of cellular processes (i.e. proliferation, differentiation, survival, apoptosis and cytokine production). Each cascade consists of a three-core member module. Each core member phosphorylates and activates the succeeding member until the final core member activates downstream kinases (e.g. transcription factors), to induce a cell response (7,8). The best studied of these is extracellular signal-regulated kinase (ERK1/ERK2) (8). Upon receptor stimulation (usually a receptor-activated tyrosine kinase), the small G-protein Ras is activated and phosphorylates the three core member cascade. The final core member, MAPK, phosphorylates proteins that regulate to cell proliferation and/or cell differentiation (7,8).

The PKA pathway can modify and influence the outcome of several signal transduction pathways, in many cells and tissues (7). In particular, in normal cells, the RI{alpha} subunit of PKA can interfere with the ERK1/2 cascade of the MAPK pathway, at the level of c-Raf-1, causing a cell-type-specific inhibition of MAPK and of cell proliferation, which has been seen in varied cell types including T- and B-lymphocytes (9–14).

Since RI{alpha} is also present in lymphocytes of CNC patients (2,15), we used these cells to examine whether MAPK is one pathway responsible for tumorigenesis in this disease. PKA activity was first examined in lymphocytes carrying the 578delTG inactivating PRKAR1A mutation (mutant, mt) and this activity was compared to that in cells obtained from unaffected relatives. As in CNC tumors, PKA activity in mutant cells was augmented; quantitative analysis indicated that most of this activity is due to the expression of type I R subunits. Isoproterenol and forskolin, which activate PKA activity in lymphocytes (16,17) were used, and the MAPK pathway assessed after stimulation with the platelet derived water-soluble phospholipid, lysophosphatidic acid (LPA, 1-acyl-sn-glycerol-3-phosphate) (18,19). Because activation of the ERK1/2 cascade of MAPK is known to lead to cell proliferation in lymphocytes and in other cell types (8,20,21), the effect of mtPRKAR1A on cell proliferation and metabolism was also studied. These studies were also performed on a tumor cell line carrying mtPRKAR1A and an in vitro construct of a mtPRKAR1A that was recently shown to be associated with increased PKA activity (22). The data suggest that indeed ERK1/2 phosphorylation is altered in mtPRKAR1A cells, a finding that may be associated with increased proliferation and metabolism and perhaps tumorigenesis.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
PKA activity
At baseline, kinase activity was higher in cells (Table 1) with the 578delTG mtPRKAR1A gene (2384±228 cpm) versus that in cells from their unaffected relatives (1290±106 cpm; P=0.02; Fig. 1A). Following exposure to cAMP, all cells responded with an increase in kinase activity (P<0.001, for all comparisons between each group's baseline and after cAMP-peak-value). However, mtPRKAR1A cells had a higher stimulation of their kinase activity (2967±265 cpm) than wild-type (wt) controls (1747±330 cpm; P=0.03). This was also true for inhibition of the cAMP-stimulated activity by a PKA-specific inhibitor (PKI): mtPRKAR1A cells maintained higher activity in response to cAMP (2214±192) than those with the wtPRKAR1A sequence (1262±95; P=0.02).


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  		Table 1. Cell lines, PRKAR1A mutations and index of studiesa
 


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  		Figure 1. Protein kinase A activity and subunit expression in mutant and normal lymphocytes. (A) PKA activity in mtPRKAR1A lymphocytes (n=8) carrying the 578delTG mutation and wtPRKAR1A cells obtained from unaffected relatives (n=3) at baseline, after exposure to cAMP, and following inhibition with a PKA-specific inhibitor (PKI): mutant cells have higher PKA activity at all stages. (B) Statistical analysis of real time amplification of the mRNA from three subunits that are expressed (PRKAR1A, PRKAR1B and PRKARCA) in the studied lymphocytic cell lines (n=11 for mutants, n=6 for unaffected relatives) using the cycle at which they were first detected (on the y-axis).

 
There was a tendency for higher total PKA activity in mtPRKAR1A cells (2.84±0.31 U/mg) versus that in controls (1.55±0.79 U/mg; P=0.091). Free PKA activity, on the other hand, was clearly higher in the former (0.57±0.1 U/mg) than the latter (0.09+0.05 U/mg; P=0.02) and, consequently, the PKA activity ratio was higher in the mutant cells (0.22±0.04 versus 0.04±0.02, respectively; P=0.03; data not shown).

Quantitative PKA subunit mRNA studies
Quantitative, real-time PCR demonstrated variable levels of the mRNAs for three of the main PKA subunits (RI{alpha}, RIß and C{alpha}) among the various cell lines, and lack of detectable expression of the remaining two (RII{alpha}, RIIß). Figure 1B shows a statistical analysis of the three subunits that were expressed in these lymphocytic cell lines. In the mtPRKAR1A cells, as expected, there was a decrease in the amount of PRKAR1A mRNA (P=0.003). These cells bear inactivating PRKAR1A mutations that lead to active degradation of the mutant message (Table 1), as we have shown elsewhere (2,4). Thus, they are expected to be haploinsufficient for RI{alpha}. There were no significant differences between mtPRKAR1A and wtPRKAR1A cells in PRKAR1B (P=0.53) or PRKACA (P=0.81) mRNA content, as this was detected by real time PCR.

ERK1/2 and proliferation assays
Basal levels of ERK1/2 and its phosphorylated form, p-ERK1/2, in all cells were assessed by immunoassay. Relatively high levels of ERK1/2 and phosphorylated ERK1/2 were seen, with no significant differences between mt- and wtPRKAR1A (P=0.1; data not shown). LPA induced ERK1/2 phosphorylation in a time- and concentration-dependent manner in both mt- and wtPRKAR1A cells without significant differences (Fig. 2). There was modest stimulation (up to 4-fold) of phosphorylation over basal levels of phosphorylated ERK1/2 at 0–30 min with 100 nM LPA (Fig. 2A). At concentrations of 0–150 nM LPA, there was no difference at 15 min in ERK1/2 phosphorylation between mt- and wtPRKAR1A cells (Fig. 2B). However, there was clearly a biphasic effect on ERK1/2 phosphorylation: at concentrations up to 150 nM there was stimulation of phosphorylated ERK1 and ERK2 in both wt- and mtPRKAR1A cells; at 200 nM LPA, there was a significant decrease in mtPRKAR1A cells (more than 75% compared with wtPRKAR1A cells; Fig. 2B). A specific MAPK inhibitor, PD98059 (50 µM) inhibited ERK1/2 phosphorylation by 85–96% in both cell types (data not shown). Viability studies in the presence of 0–100 µM PD98059 showed no effect of the inhibitor on cell viability (data not shown).



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  		Figure 2. Time–concentration curves of LPA-induced phosphorylation of ERK1/2. Cells were stimulated with 100 nM LPA for 0–30 minutes at 37°C (A) or stimulated with increasing concentrations (0–100 nM) of LPA for 15 min (B). Cells were lysed and gel electrophoresis and immunoblot assays were performed as described in the Methods. In (A) and (B), levels of phosphorylated ERK1/2 are given as fold-stimulation of ERK1/2 over control (basal) ERK1/2, and as relative band densities in arbitrary units, respectively. In (A), control values are given as 1-fold stimulation. In (B) control values are given as a relative density of 100. Results are mean±SEM of three experiments per each point.

 
In summary, these data confirmed the presence of the ERK1/2 cascade in the investigated cell lines and suggested that this cellular pathway functions in a similar manner in normal and mutant cells, with the exception of the response to higher doses of LPA (see above).

When forskolin and isoproterenol (30 µM) were tested in LPA-stimulated cells, significant differences in phosphorylated ERK1/2 content between normal and mutant cells were seen. In the former, both PKA stimulants inhibited LPA-induced phosphorylation, while phosphorylation in the latter was enhanced (Fig. 3). Increasing concentrations of forskolin (Fig. 4A) and isoproterenol (Fig. 4B) caused a similar effect on LPA-induced (100 nM) phosphorylation, with inhibition and stimulation occurring, respectively, in normal and mutant cells. However, the effect of isoproterenol was greater (up to 3-fold in normal cells and 10-fold in mutant cells) than forskolin. Non-LPA-stimulated (control) phosphorylated ERK1/2 was also affected in the same manner by isoproterenol and forskolin (data not shown). Again, a biphasic effect was seen in all experiments (Figs 3 and 4).



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  		Figure 3. ERK1/2 phosphorylation induced by increasing concentrations of LPA; effect of forskolin and isoproterenol. Cells were pre-incubated for 10 min with 30 µM forskolin (A), or isoproterenol (B), followed by incubation for 15 min with increasing concentrations of LPA (0–200 nM). Cells were lysed and protein was separated by gel electrophoresis and analyzed by immunoblot assay as described in the Methods. FSK, forskolin; Iso, isoproterenol; LPA, lysophosphatidic acid. Values are presented as relative density, percentage of LPA-induced control stimulation. In (A) and (B), control values are given as a relative density of 100. Results are representative of three experiments for each point.

 


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  		Figure 4. Effect of increasing concentration of forskolin and isoproterenol on LPA-induced ERK1/2 phosphorylation. Cells were pre-incubated for 10 minutes with increasing concentrations (0–100 µM) of forskolin (A) or isoproterenol (B) and stimulated for a further 15 min with 100 nM LPA. Cells were lysed and protein was separated by gel electrophoresis and analyzed by immunoblot assay as described in the Methods. Values are presented as relative band density, percentage LPA-induced control stimulation. In (A) and (B), control values are given as a relative density of 100. Results are representative of three experiments for each point.

 
To determine if differences exist in growth patterns between normal and mutant cells, cell number and percent cell viability were determined at regular intervals for a total of 4 days (Fig. 5). From 0 to day 1 there was an overlap in basal growth rate between normal and mutant cells; however, from day 1 to day 4, mutant cell growth rate surpassed that of normal cells by 2–3-fold. In the presence of 100 nM LPA, the growth of both cell types increased; however, the growth rate in mutant cells was greater than that in normal cells by 1.8–2.5-fold (Fig. 5A; P<0.05). Cell viability remained constant in both cell types, at approximately 90% (data not shown). In the presence of forskolin (30 µM), as in the MAPK studies above, there was an inhibition of LPA-induced cell growth in normal cells and stimulation of growth in mutant cells (Fig. 5B).



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  		Figure 5. Cell proliferation in normal and mutant cells: effect of forskolin. Cells, plated at 1.5x105 cells/ml were incubated in the presence and absence of 100 nM LPA for a four day period (A), or pre-incubated with 30 µM forskolin for 10 min, followed by incubation with 100 nM LPA for 4 days (B). Cell count was assessed at one-day intervals. Results in (A) and (B) are expressed as cell number (1x104) or as cell number, percentage of control (basal) cell growth, respectively. Results reflect the mean±SEM of three experiments.

 
In addition, an indirect measure of cell proliferation, cell metabolism, was examined to determine whether these data would agree with our studies on cell proliferation (Fig. 6). In cell titer 96 AQ assays, cell metabolism was increased in a dose-dependent manner by LPA, and correlated (one-to-one) with cell number (data not shown). In the presence of forskolin (30 µM), LPA-induced metabolism was inhibited in normal cells (Fig. 6A and C) and stimulated in mutant cells (Fig. 6B and C). The effect of forskolin on cell metabolism induced by increasing concentrations of LPA was biphasic for both normal and mutant cells. Thus, indeed, this assay correlated well with the dose–response biphasic effect on ERK1/2 phosphorylation that was seen in the MAPK studies (see above) and followed the changes detected between mutant and normal cells after their stimulation with forskolin and isoproterenol.



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  		Figure 6. Effect of forskolin on LPA-induced cell metabolism. Normal (A) and mutant (B) cells were plated at 5x103 cells/ml in 96-well plates and equilibrated at 37°C for 1–3 h before the addition of 30 µM forskolin, followed by incubation with 0–200 nM LPA for 24–36 h. Cell titer 96 AQ solution was added for an additional 4 h. Plates were read by an ELISA reader at 460 nm. Results in (A) and (B) are expressed as percentage of LPA-induced control absorbance at 0 nM LPA. Results in (C) are expressed as percentage of control of absorbance at 0–100 nM LPA. In (A)–(C), control values are given as an absorbance of 100% of basal values. Values are representative of six experiments, and are the mean±SEM of five wells per experimental point.

 
ERK1/2 phosphorylation and proliferation assay in a pituitary tumor cell line
To validate our data on mtPRKAR1A lymphocytes, MAPK (ERK1/2) activity and cell proliferation were also assayed in primary cell lines established from tissues of patients with a wtPRKAR1A gene [skin and adrenal (Fig. 7A and B), cell line data 1 and 2, respectively] and from a pituitary tumor (cell line 3) bearing the mtPRKAR1A gene (c.578delTG) in the hemizygous state (data not shown) in the presence and absence of forskolin. Phosphorylated ERK1/2 was decreased in normal fibroblasts by 30–60%, while activity in mutant cells was enhanced to over 40% (P<0.001; Fig. 7A). Cell metabolism (an indirect measure of proliferation) was affected in a manner analogous to that seen in lymphocytes (Fig. 5) bearing the c.578delTG and other PRKAR1A-inactivating mutations (Table 1); there was a 20–25% inhibition in wtPRKAR1A cells, whereas in mtPRKAR1A cells metabolism was stimulated by 30% in response to forskolin (Fig. 7B).



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  		Figure 7. Analysis of primary and/or transfected cell lines. (A, B) Primary cell lines from normal tissues of two different subjects with wtPRKAR1A (1 and 2), and a pituitary tumor from a CNC patient bearing the c.578del TG PRKAR1A (cell line 3) were assayed for p-ERK1/2 activity in response to forskolin (A) and for metabolic activity (reflecting to some extent proliferation) (B), as described in the Methods. Results in (A) are relative band density, percentage of control stimulation and represent the mean±SEM of four experiments. Values in (B) are percentage of control absorbance and are mean±SEM of seven experiments using five wells per experimental point. (C) Forskolin responses of p-ERK1 (left) and p-ERK2 (right) in COS-7 cells transfected with the RI{alpha} {Delta}184–236 construct or the vector (pREP4), as described by Groussin et al. (23).

 
ERK1/2 phosphorylation in COS-7 cells bearing the RI{alpha} {Delta}184–236 mutation
In COS-7 cells transfected with a PRKAR1A construct bearing the RI{alpha} {Delta}184–236 mutation which leads to increased PKA function (22), forskolin at 10 µM increased the presence of phosphorylated ERK1 and ERK2 activity by 59 and 141%, respectively versus control. Phosphorylated ERK1 and ERK2 were also greatly increased over the plain vector-transfected cells by 38 and 74%, respectively (Fig. 7C).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
In the present study, the presence of PKA activity in lymphocytes carrying a PRKAR1A germline mutation (c.578delTG) and in normal controls was first confirmed. It was then shown that most of this PKA activity depends on the expression of type I PKA regulatory subunits, namely RI{alpha} and RIß (Fig. 1). The PKA activity of mtPRKAR1A cells was greater than that of wtPRKAR1A cells, suggesting that the RI{alpha} decrease and the resulting disturbance of the balance between the two type I regulatory subunits cause substantial differences in cAMP signaling in these cells. The data on PKA activity are similar to what we have seen previously in CNC tumors (carrying the same mutation) (4); in these tissues it was postulated that type II PKA activity was responsible for the increased responses to cAMP (3).

Increased PKA activity in lymphocytes carrying the c.578delTG PRKAR1A inactivating mutation indicated that these cells, despite being only haploinsufficient for PRKAR1A, do have abnormalities of the cAMP-dependent PKA signaling system. Thus, these cells are perhaps a good system for studying signaling alterations that are associated with mtPRKAR1A. Indeed, we used these lymphocytes to study the interactions of PKA with a major growth and proliferation control pathway, the ERK1/2 cascade of the MAPK signaling system. Several lymphocytic cell lines with PRKAR1A mutations that have the same molecular effect on PRKAR1A (absence of the mutant transcript) due to nonsense-medicated RNA decay (2) were used for these experiments (Table 1).

A dose-dependent effect of PKA activation on MAPK activity, as measured by phosphorylated ERK1/2 induction, was found in these cells; it was also found that this effect correlates well with indices of cell proliferation and metabolism. Although ERK and phosphorylated ERK1/2 activity were similar in both mutant and normal cells after the application of LPA, it was found that phosphorylated ERK1/2, cellular proliferation and cell metabolism were affected in opposite ways in mt- versus wtPRKAR1A lymphocytes. The high basal levels of ERK1/2 and phosphorylated ERK1/2 in both normal and mutant cells were not unexpected in EBV-transformed B-lymphocytes (23,53), which have been shown to contain a constitutively active Jak/STAT pathway (24). In the present studies and those of others (18), LPA was shown to activate the ERK1/2 cascade of the MAPK pathway in EBV-transformed B-lymphocytes. In our study, LPA induced only a modest stimulation of phosphorylated ERK1/2 activity, which was similar in both mutant and normal cells for LPA concentrations up to 150 nM (Fig. 2). The possibility exists that this similar effect may reflect the high basal levels of ERK1/2 and phosphorylated ERK1/2, which could mask any difference in stimulation between normal and mutant cells. On treatment of cultures with 50 µM PD98059, the MEK1 inhibitor of the ERK1/2 cascade, followed by treatment with 100 nM LPA, MAPK activity was almost completely abolished in both cell types (data not shown). However, studies with increasing PD98059 concentrations (0–100 µM, using Trypan blue dye exclusion) showed no effect of the inhibitor on cell viability (data not shown). These experiments, as well as those of Rosskoft et al. (21), strongly support the idea that LPA stimulates the ERK1/2 cascade of the MAPK pathway in EBV-transformed B-lymphocytes, and that this pathway functions similarly in both wt- and mtPRKAR1A cells.

The interaction of the PKA and the MAPK pathways, previously seen in T- and B-lymphocytes and in other cell types (9,12–14), is also shown in the present studies. Interestingly, however, although an inhibition of ERK1/2 phosphorylation is seen in normal cells, a profound difference occurs in the effect of PKA in mutant cell lines (Figs 3 and 4). In mutant cells, PKA caused an enhanced stimulation of LPA-induced phosphorylation. In normal cells, the inhibitory effect of PKA has been attributed to an inhibition of C-Raf-1 activity in the ERK1/2 cascade, by type I PKA (25). This inhibition is thought to occur through the phosphorylation of Ser259 with modulatory phosphorylation of Ser43 and Ser621, of the C-Raf-1 molecule (26). However, the Raf isoforms are differentially sensitive to PKA. In cells having both the B and C forms of Raf, PKA can inhibit C-Raf and stimulate B-Raf to inhibit and stimulate cell proliferation though ERK1/2, respectively (27,28). The action of PKA on B-Raf is, in some cell types, through the activation of the small G-protein, Rap-1 (29–31). In normal cells, a balance exists between stimulation and inhibition of ERK1/2 activity by PKA, with inhibition being more dominant. This dominance appears to be due to the presence of a greater number of total type I PKA (70–80%) and a smaller number of type II PKA (20–25%) molecules in lymphocytes (29) and a higher affinity of cAMP for the RI{alpha} subunit (6,32). In the presence of RI{alpha}-haploinsufficiency, as in CNC-affected lymphocytes (4), the balance may shift towards activation of ERK1/2 through another PKA subunit, perhaps the RIß subunit, in CNC cells. The underlying mechanism may be, as seen in other systems (32), a biochemical compensation or substitution of one subunit for the other (e.g. RIß for the RI{alpha}), which may lead to cell proliferation instead of inhibition (27). B-Raf regulation by PKA may be responsible for this phenotype (33). Indeed, we have shown in CNC and normal lymphocytes the presence of both the B and C isoforms of Raf (unpublished data). Accordingly, a decreased or dysfunctional RI{alpha} molecule has been implicated in tumorigenic cell proliferation (34), and other investigators have shown that normal RI{alpha} activity is needed for the cAMP-mediated inhibition of proliferation of T- and B-lymphocytes (25,35).

The effect of isoproterenol on ERK1/2 phosphorylation was 3- and 10-fold greater than that of forskolin in normal and mutant cells, respectively. This difference could reflect a difference in receptor- versus non-receptor-mediated cAMP stimulation of the PKA holoenzyme. Since multiple intracellular pathways can be affected by cAMP independently of PKA (17,31,36), the data may indicate an interference by other cAMP-related pathways on the overall PKA/MAPK interaction (31,36).

Our cell proliferation and metabolism studies indicated a clear difference between normal and mutant cells (Figs 5 and 6); furthermore, the forskolin and isoproterenol effects on these indices followed the previously described patterns of ERK1/2 phosphorylation. Indeed, it has previously been shown that RI{alpha} mediates the PKA inhibitory effects on T-lymphocyte proliferation (29). The biphasic effect of LPA and the PKA stimulants that was seen in both MAPK and cell metabolism experiments is not unique to LPA, since it has been shown with other MAPK stimulants (37–39) and cell proliferation studies (37) in several cell systems. A similar biphasic effect was seen with both forskolin and isoproterenol (Fig. 4). It is possible that, at higher agonist concentrations, the biphasic effects could represent a saturation of pathway receptor (MAPK and PKA) and enzyme (adenyl cyclase, MAPK phosphatases) kinetics (40,41).

These studies demonstrated that PRKAR1A differentially affects ERK1/2 and indices of cell proliferation and metabolism in wt- and mtPRKAR1A lymphocytes. We have confirmed these data in at least one tumor cell line, i.e. in a cell line established from a pituitary adenoma of a patient with CNC (4,42). Indeed, our assays of phosphorylated ERK1/2 and metabolism/proliferation on this primary cell line carrying the c.578delTG PRKAR1A mutations (Fig. 7A and B) were surprisingly similar to the data in the lymphocytes carrying the same and other PRKAR1A-inactivating mutations. Finally, an in vitro construct of a PRKAR1A mutation (R1{alpha} {Delta}184–236) that leads to higher levels of PKA activity in COS-7 cells (and unlike other PRKAR1A mutations does not lead to a nonsense transcript) (23) had similar effects on phosphorylated ERK1/2 levels (Fig. 7C). These data collectively suggest that tumorigenesis in CNC may indeed involve MAPK-signaling pathways.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Cell lines
Lymphocytes were obtained from patients with CNC, their unaffected relatives and other normal controls using the Ficol® method [obtained from Amersham Pharmacia (Piscataway, NJ, USA)] under a research protocol approved by the Institutional Review Board of the National Institute of Child Health and Human Development. They were transformed by Epstein–Barr virus (EBV) and maintained in culture by standard methods (23). These methods and western blotting for PRKAR1A protein, as well as the families of the individuals and their PRKAR1A mutation status have been described in detail elsewhere (2,4). Only cell lines in which mtPRKAR1A undergoes nonsense mRNA-mediated decay (NMD) (2,4) were chosen for the present investigation. Table 1 summarizes the cell lines, mutation status and the assays they were used in, for the purposes of the present study. The pituitary tumor (4) and control primary cell lines were established by standard methods. COS-7 cells and their transfections, constructs, and vector information were published recently (22).

PKA activity determinations
PKA activity was measured, as described previously (4), using [{gamma}-32P] dATP (deoxyadenosine 5'-[{gamma}-32P]-triphosphate, Amersham Pharmacia Biotech, NJ, USA), in cell extracts that had been snap-frozen in liquid nitrogen. Total kinase activity represents enzymatic activity after stimulation with cAMP; total PKA-specific activity represents the difference between PKA activity before and after the addition of protein kinase inhibitor (PKI). A ratio was calculated after the determination of free PKA activity, according to the following formula:

All determinations of PKA activity were done twice for each cell line, corrected for protein content, and an average value was calculated for each experiment.

The data from all cells shown in Figure 1 were compared with STATISTICA software (StatSoft Inc., Cary, NC, USA) using the t-test for individual comparisons between the two diagnostic groups [cells with the 578delTG mtPRKAR1A (n=8) and cells from unaffected first-degree relatives with wtPRKAR1A (n=3)]. All data are shown with the mean±standard error of the mean (SEM). A P-value of less than 0.05 was considered to indicate significance; a P-value of less than 0.1 was interpreted as showing a tendency towards a significant difference.

Real time quantification of mRNA
Real-time polymerase chain reaction (PCR) was performed using the Light Cycler apparatus (Roche). PCR reactions were performed in a 20 µl final volume, probe 0.5 µM, MgCl2 3.5 mM according to the manufacturer's recommendations. Sequences used for the PRKAR1A, PRKAR1B, PRKAR2A, PRKAR2B and PRKACA probes were obtained from the genome databases. All results were normalized against a housekeeping gene expression (GAPDH). A melting curve and the cycle at detection (shown in Fig. 2A–C) were analysed with the software of the Light Cycler apparatus (43). For all statistical analyses, the cycle at which each expressed subunit was detected was used. We then employed t-test using the STATISTICA software, as described above. A total of 11 mtPRKAR1A cell lines (carrying inactivating PRKAR1A mutations that lead to NMD) and six controls were used for this analysis (Table 1).

Materials for cultures, growth, proliferation and MAPK assays
Materials were obtained from the following sources: RPMI 1640 media, AIM V media, Dulbecco's Modified Eagles Media (DMEM), fetal bovine serum (FBS), Hepes, antibiotics/antimycotics and Trypan blue were from GIBCO-BRL Life Tech. (Rockville, MD, USA); forskolin, L-{alpha}-lysophosphatidic acid, isoproterenol, phenylmethaneosulfonyl fluoride (PMSF), Triton-X 100, horse donor herd serum and Ponceau Reagent were from Sigma-Aldrich (St Louis, MO, USA); leupeptin, aprotinin and 4-nitrophenylphosphate (4-Npp) were from Roche Molecular Biochem (Indianapolis, IN, USA); PD98059 was from CalBiochem (San Diego, CA, USA); phospho-anti ERK1/2 and anti-ERK1/2 were from Cell Signaling (Beverly, MA, USA); anti-mouse IgG was from Oncogene Research Products (Darmstadt, Germany); anti-B lymphocyte CD23 and Invision System staining kit were from DAKO (Carpenteria, CA, USA); anti ß-actin was from Santa Cruz Biotechnology (Santa Cruz, CA, USA); BCA protein assay kit from Pierce (Rockford, IL, USA); bovine serum albumin fraction V was from ICN (Irvine, CA, USA); 10x Tris/glycine/SDS buffer, 10x Tris/glycine buffer, blotting-grade non-fat dried milk, and Tween 20 were from BioRad (Hercules, CA, USA); Protran nitrocellulose membranes were from Schleicher and Schuell (Keene, NH, USA); Novex 10% Tris/glycine gels were from Invitrogen (Carlsbad, CA, USA); cell titer 96 AQ Assay was from Promega Corp. (Madison, WI, USA); and enhanced chemiluminesence (ECL) blotting detection reagent and Ficol were from Amersham Pharmacia (Piscataway, NJ, USA).

Cell cultures and plating
Lymphocytes were maintained as cell suspensions in RPMI 1640 media with L-glutamine, 10% FBS and 1% antibiotic and antimycotic agents; media was changed every 3 days. The MAPK, cell growth and proliferation studies were conducted on five CNC-affected mutant cell lines and on 11 normal cell lines (Table 1), in log growth phase. The B-lymphocytic characteristics of each cell culture were assessed by staining with antibodies to the B-cell surface antigen, CD23 (44). This antigen is primarily expressed on the surface of B-lymphocytes and monocytes (45,46) and is strongly expressed on EBV-transformed B-lymphocytes (47). All cell lines expressed the CD23 cell surface antigen (data not shown). The immunological characteristics of EBV-transformed lymphocytes have been discussed elsewhere (48).

Primary cell lines were maintained in Dulbecco's modified Eagles medium with 15% FBS, 1% horse donor herd serum and 1% antibiotic/antimycotic agents. The medium was changed every 5–7 days. COS-7 cells were maintained in similar media without horse-donor herd serum and treated as described elsewhere (22).

For immunoassays, lymphocytic cultures were washed, resuspended and diluted to 1–2.5x106 cells/ml in PBS (pH 7.4), and transferred to 1.5 ml Eppendorf tubes before the addition of PKA (forskolin and isoproterenol) and MAPK (LPA) stimulants. Since serum contains high levels of BSA-bound LPA, in growth studies, cells were plated in 24-well cluster plates at 5x104 cells/well, in AIM V serum-free media plus 1% antibiotics, followed by the addition of stimulants. Likewise, in cell metabolism studies, lymphocytic cell lines in AIM V serum-free media were plated in 96-well cluster plates at 5x103 cells/well, and incubated for 1–3 h, before the addition of stimulants for 18–36 h.

For immunoassays of the primary cultures and COS-7 cells, all cells were plated at a density of 3–7x104 cells/ml in 35 mm culture dishes and 12-well plates, respectively, and allowed to attach for 12–18 h at 37°C. Cells were then incubated with and without forskolin for a further 30 min (primary cultures) and 6 h (COS-7 cells). For cell metabolism studies, cells were plated at 1x103 cells/well in 96-well culture plates, and incubated at 37°C for 12–18 h, followed by incubation for 36–48 h in media with and without forskolin.

Electrophoretic separation and immunoblotting assays
The following procedures are a modification of the methods of Hirasawa et al. (49). Cell lysates from lymphocytic cell lines were obtained when stimulated cells were placed on ice immediately after incubation, and centrifuged (1000g for 20 s). Cell pellets were resuspended in lysis buffer (pH 7.3) containing 20 mM Hepes, 10% glycerol, 1% Triton X-100, 50 mM NaF, 1 mM NavO4 and protease inhibitors (0.02 mg/ml aprotinin, 0.02 mg/ml leupeptin, 1 mM PMSF and 2.5 mM 4-Npp), and were homogenized at 4°C. Protein assays were performed on the complete cell lysate. Cell lysates from primary cell lines or transfected COS-7 cells were obtained when cells after stimulation were immediately placed on ice, washed with ice-cold PBS and lysis buffer was added (as described above). Cells were scraped into Eppendorf tubes, and lysates were left on ice for 30 min to allow further cell lysis. All lysates were diluted to 1–2.5 µg/ml protein. To assure uniformity of loading, equal amounts of protein were added per well, as ascertained by Ponceau staining, or by antibody to B-actin. Samples were subjected to SDS–PAGE and transferred to nitrocellulose membranes. Membranes were blocked for 1 h at room temperature with 7% non-fat dried milk in Tris buffered saline (with 1% Tween 20, or TBST).

MAP kinase and phosphorylated proteins were detected by immunoblot technique using polyclonal p44/p42 antibody (ERK1/2) raised in rabbit and monoclonal phosphorylated p44/p42 (p-ERK1/2, phosphorylated at Thr202 and Try204, i.e. activating phosphorylations) raised in mice, respectively, as primary antibodies, diluted in 5% BSA in TBST. Horseradish peroxidase-conjugated antibody, against mouse and rabbit IgG, diluted in 7% non-fat dried milk in TBST, was used as secondary antibodies. Bands were detected by ECL reagent and quantitated by densitometer scanning (Molecular Dynamics, Sunny Vale, CA, USA).

Cell proliferation and metabolism measurements
Cell proliferation was determined using direct cell counts by hemocytometer, and by the cell titer 96 AQ assay. To measure cell proliferation by direct cell count, cells were plated as stated above, and counted each day for 4 days using Trypan blue cell viability solution. The cell titer 96 AQ assay, a colorimetric assay, measures cell ‘metabolism’ in a way that directly correlates (one–one) with cell number (50–53). It measures the conversion of MTS tetrazoleum salt to formazan by cellular dehydrogenase activity. Following cell stimulation, a 20 µl aliquot of cell titer 96 AQ solution was added to cultures for a 4-h incubation period at 37°C. Absorbance was determined at 490 nm, in an ELISA plate reader (Biorad model 550).

Statistical analyses for the MAPK and proliferation assays
All data are shown with the mean±SEM (Figs 2–7). For these assays, the data were analyzed by ANOVA followed by Turkey's multiple comparisons test. A P-value of less than 0.05 was considered significant in all experiments.


   ACKNOWLEDGEMENTS
 
We thank Dr Michael A. Beaven (NHLBI, NIH) for his advice regarding methodology and for a critical review of the manuscript; we also thank Dr Y.S. Cho-Chung for her help in measuring PKA activity in various cells and tissues and for guidance in the protein kinase A field. We would like to acknowledge partial support of this study by the ‘GIS-INSERM Institut des Maladies Rares’ (to J.B.). Finally, we thank the patients that participated in the Carney complex NICHD study (95-CH-0059) and donated their samples for the establishment of the studied cell lines.


   FOOTNOTES
 
* To whom correspondence should be addressed at: Section on Endocrinology and Genetics, DEB, NICHD, NIH, Building 10, Room 10N262, 10 Center Dr. MSC1862, Bethesda, MD 20892-1862, USA. Tel: +1 3014964686; Fax: +1 3014020574; Email: stratakc{at}mail.nih.gov Back


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
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