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INSERM, U-344, Molecular Endocrinology, Faculté de Médecine Necker (S.B., S.K., S.J., M.L., P.A.K., V.G.), 75730 Paris, France; Laboratory of Molecular Biology and Genetic Engineering, University of Liege (S.K., J.A.M.), 4000 Sart-Tilman, Belgium; Laboratory of Molecular Biophysics, Institut de Biologie Structurale, Commissariat à l’énergie atomique-Centre National de la Recherche Scientifique (D.M.), 38027 Grenoble, France
Address all correspondence and requests for reprints to: Dr. Vincent Goffin, INSERM, U-344, Molecular Endocrinology, Faculté de Médecine Necker, 156 rue de Vaugirard, 75730 Paris Cedex 15, France. E-mail: goffin{at}necker.fr
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Introduction |
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PRL exists in several molecular isoforms resulting from various posttranslational modifications, including proteolytic cleavages, glycosylation, and phosphorylation (for a review, see Ref. 12). The occurrence in vivo of PRL phosphorylation has been clearly demonstrated in bovine pituitary (13), where serine 90 was identified as the major phosphorylation site of bovine PRL (14). In other species the in vivo demonstration of PRL phosphorylation has been less definitive (15, 16), and the physiological relevance of phosphorylation sites identified by in vitro incorporation of 32P into PRL (17, 18) remains questionable. In humans, although pituitary-purified human PRL (hPRL) preparations obtained from the NIDDK were shown to contain traces of phosphorylated forms (19, 20), definite identification of phosphorylated amino acid(s) awaits further investigation.
Several reports have shown that phosphorylation alters the biological properties of PRL. For example, phosphorylated bovine PRL displays reduced mitogenic activity in the Nb2 cell proliferation bioassay compared with the nonphosphorylated isoform (21). Accordingly, dephosphorylation of hPRL slightly increases its bioactivity (19, 20). Still more remarkably, phosphorylation of serine 177 confers antagonistic properties to rat PRL (rPRL) in the same Nb2 cell proliferation assay (18, 19). This serine residue is highly conserved among PRLs from different species (22) and is homologous to serine 179 in hPRL. Based on the antagonistic properties demonstrated for phosphorylated rPRL, a molecular mimic of a putative S179-phosphorylated hPRL has been engineered in Walker’s laboratory by substituting an aspartate for the wild-type (WT) serine 179, generating the so-called S179D-hPRL analog (20). In a first report these researchers reported that S179D-hPRL strongly antagonizes the mitogenic activity of hPRL on Nb2 cells, thereby extrapolating to the human hormone the functional consequences of Ser177 phosphorylation earlier reported for rat PRL (20). Surprisingly, contrary to all previous reports describing the mechanism of action of PRL or GH antagonists (23, 24, 25, 26, 27), S179D-hPRL appeared to antagonize hPRL effects through a noncompetitive inhibition of receptor activation (20). This encouraged Walker and colleagues to investigate the molecular basis of this unusual, although apparently highly potent, mechanism of antagonism. Using the immunoblot approach to investigate activation of the JAK2/STAT5 cascade, these researchers showed that S179D-hPRL strongly activates STAT5 while minimally activating JAK2 (28). Based on these observations, they suggested the involvement of a kinase other than JAK2 in STAT5 activation, and hence, they hypothesized that the antagonistic activity of S179D-hPRL may result from activation of signaling molecules/pathways other than those known to be involved in hPRL signaling (1) and still to be identified.
For many years the development of hPRL antagonists that could be of use in reducing breast cancer progression has been one of our goals (8, 29). In this perspective we have previously engineered several hPRL analogs, one of which (G129R-hPRL) appeared to display competitive antagonistic properties in various PRLR-mediated bioassays (25, 26), including hPRL-induced signaling and proliferation of breast cancer cells (27). However, due to its lower affinity, the analog had to be used at a significant molar excess vs. WT hPRL (1:10 to 1:50 ratio) to exhibit significant antagonism, which justified further efforts in developing more potent hPRL antagonists. In this respect the conclusions claimed by Walker et al. in their two recent reports (20, 28) appeared of great interest and prompted us to take further advantage of the antagonistic properties of S179D-hPRL. Thus, the present work was initiated with the aim to engineer more potent hPRL antagonist by combining the two point mutations [S179D (20) and G129R (25)] individually conferring antagonism to hPRL analogs. However, we failed to confirm the conclusions of Walker’s articles regarding S179D-hPRL properties, as this mutant clearly acts as an agonist, and not an antagonist, on the PRLR. The goal of the present report is to better characterize the biological properties of this analog in various cell bioassays and to discuss why antagonistic properties were erroneously attributed to S179D-hPRL.
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Materials and Methods |
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Hormones
All of the hormones used in this study were produced by
recombinant technology: WT hPRL (30), the binding site 2
analog G129R-hPRL (Gly129 replaced with Arg)
(31), and the molecular mimic of phosphorylated hPRL,
so-called S179D-hPRL (Ser179 replaced with Asp).
The mutants were produced in Escherichia coli BL21(DE3)
using the pT7L expression vector (32), then purified as
described previously (25, 31, 32, 33).
Cells
Rat Nb2 lymphoma cells were obtained from P. W. Gout
(Vancouver, Canada). As previously reported (26), HL5
cells are a clone of 293 fibroblastic cells stably transfected with the
plasmids encoding the hPRLR (34) and a PRL-responsive
reporter gene. The latter contains the sequence encoding the luciferase
gene under the control of a six-repeat sequence of the lactogenic
hormone response element (LHRE; which is the DNA-binding element of
STAT5), followed by the minimal thymidine kinase promoter. T-47D cells
(human mammary tumor epithelial cell line) were provided by M. N.
Norman (Bristol, UK).
Site-directed mutagenesis
Construction of the S179D-hPRL mutated cDNA was performed by the
oligonucleotide-directed mutagenesis method using the Chameleon double
stranded, site-directed mutagenesis kit from Stratagene
(La Jolla, CA), following the manufacturer’s instructions. The
pT7L-hPRL expression vector (32) was used as a template
for mutagenesis. The sequence of the mutated oligonucleotide was the
following (5'
3' noncoding strand, mutated codon
underlined): GTC GAT TTT ATG GTC ATC CCT GCG
TAG. The selection primer (XmnI restriction site; see
manufacturer’s instructions) was the following: 5'-CAT CAT TGG AAA ACG
CTC TTC GGG GCG-3'. Clones containing the expected mutation were
identified by DNA sequencing on the pT7L-hPRL plasmid
(32).
Biochemical properties
Production and purification of proteins. Recombinant WT hPRL
and hPRL analogs were overexpressed in a 1-liter culture of E.
coli BL21(DE3) and purified as previously described
(32). Briefly, when the OD600 of
bacterial cultures reached 0.7–0.9, overexpression was induced using 1
mM isopropylthiolgalactoside (IPTG) for 2
(20) or 4 h (32)
(OD600, 
2.5 after 4 h). Cell lysis was
performed using a cell disintegrator (Basic Z, Cell D,
Roquemaure, France). Proteins were overexpressed as insoluble
inclusion bodies that were solubilized in 8 M
urea (5 min at 55 C, then 2 h at room temperature) and refolded by
continuous dialysis (72 h, 4 C) against 50 mM
NH4HCO3, pH 8. Proteins
collected at this step of the protocol were referred to as nonpurified.
Alternatively, refolded proteins were concentrated by tangential flow
ultrafiltration (500 ml/min flow rate) using a YM10 Miniplate
bioconcentrator (Millipore Corp.-Amicon, Bedford, MA).
After centrifugation (10 min, 9000 x g) of
concentrated protein, supernatants were purified by gel filtration
chromatography using a high resolution Sephacryl S-200 column
(Amersham Pharmacia Biotech) equilibrated in 50
mM
NH4HCO3 and 150
mM NaCl, pH 8. Fractions corresponding to
monomeric hPRL (WT and analogs) were collected, pooled, quantified by
the Bradford method (Bio-Rad Laboratories, Inc.,
Ivry-sur-Seine, France), aliquoted, and stored at -20 C. This
final fraction was referred to as purified.
SDS-PAGE and protein quantification. Protein size and purity were assessed using 15% SDS-PAGE under reducing (ß-mercaptoethanol) or nonreducing conditions (35). Gels were stained using Coomassie blue. Protein preparations were quantified by Bradford protein assay, using BSA as the reference.
Circular dichroism spectroscopy. A Jobin Yvon CD6 circular dichroism spectropolarimeter with a thermostated sample holder was used. Data were recorded at 25 C using 0.1-cm cells with an interval of 1 nm and an integration time of 5 sec. The sample concentration was 0.4 mg/ml in 25 mM NH4HCO3 and 150 mM NaCl, pH 8. Each spectrum is an average of three scans, corrected for buffer. Helicity was calculated as described (36).
Apparent molecular mass. The apparent molecular mass (MM) of WT hPRL and hPRL analogs was estimated using analytical gel filtration chromatography (HR S-200) according to calibration from several MM markers: dextran blue (void volume), BSA (68 kDa), carbonic anhydrase (30 kDa), and myoglobin (17.5 kDa).
Isoelectric focusing. The isoelectric point (pI) of hPRL (WT and mutants) was determined by isoelectrofocusing. Electrophoresis was performed under continuous cooling; electrode solutions were 20 mM acetic acid and 20 mM NaOH. The gel contained polyacrylamide (5.5%), glycerol (10%), and ampholytes (5.5%) in a range of pH 3–10. A prerun of 15 min (200 V) was applied to the gel before sample loading to allow formation of the pH gradient. Ten micrograms of protein diluted in sample buffer (5.5% ampholytes and 10% glycerol) were loaded on the gel, then the run was performed at 200 V until the visible band corresponding to methyl red (pI 3.75) was focused. Gels were fixed in 20% trichloroacetic acid, then in 40% ethanol/10% acetic acid. Staining was performed using Coomassie blue and destaining in 40% ethanol/10% acetic acid. The pI of hPRL samples were estimated by comparison with pI markers (Amersham Pharmacia Biotech).
Biological properties
Binding studies. Binding affinity of S179D-hPRL was
determined using cell homogenates of the HL5 clone (expressing the
human PRLR), following the procedure previously described
(26). Briefly, hPRL was iodinated using Iodogen
(33), and its specific activity was in the range of 40–50
µCi/µg. Binding assays were performed overnight at room temperature
using 150–300 µg cell homogenate protein in the presence of 30,000
cpm [125I]hPRL and increasing concentrations of
unlabeled WT hPRL or S179D-hPRL. Results are representative of three
independent experiments performed in duplicate. The relative binding
affinity of the mutant was calculated as the ratio of its
IC50 with respect to that of WT hPRL.
In vitro cell bioassays.
Nb2 cell proliferation assay.
The reference bioassay
for lactogenic hormones is the lactogen-induced proliferation of rat
Nb2 lymphoma cells (37, 38). The Nb2 cell line was
routinely maintained in RPMI 1640 supplemented with 10% horse serum,
10% heat-inactivated FCS, 2 mM glutamine, 50
U/ml penicillin, 50 µg/ml streptomycin, and 100
µM ß-mercaptoethanol. The proliferation assay
was performed as initially described (37) with minor
modifications. Briefly, the assay was performed in 96-well plates using
2 x 104 cells/well on a starting day in a
final volume of 200 µl, including hormones. As it will be explained
in Results, lower cell densities were also tested according
to the protocol of Chen et al. (20). Cell
proliferation was estimated after 3 d of hormonal stimulation by
adding 10 µl WST-1 tetrazolium salt (Roche, Meylan,
France) as previously described (39). This survival
reagent is metabolized by mitochondria of living cells, which leads to
an increase in the OD measured at 450 nm (OD450)
in a manner that is proportional to the number of cells counted by
hemocytometer (data not shown). A minimum of 30 min was required to
get homogenous colorimetric reaction, then OD450
values increased for the various experimental conditions proportionally
to the time of incubation with WST-1. The linearity of
OD450/cell density ratios up to absorbance values
of 3–3.5 was confirmed by performing preliminary cell dilution
experiments (not shown). We report data obtained after 2–4 h of
reaction (which fall within this absorbance range) to allow detection
of minimal cell proliferation under low stimulation conditions.
Antagonistic properties of hPRL mutants were assessed in competition
assays using 200 pg/ml or 1 ng/ml WT hPRL. The experiments were
performed at least three times in triplicate or quadruplicate.
Human PRLR transcriptional bioassay.
The HL5 clone (human
PRLR-LHRE-luciferase, clone 5) (26) was routinely cultured
in DMEM-Nut F12 medium supplemented with 10% FCS, 2
mM glutamine, 50 U/ml penicillin, 50 µg/ml
streptomycin, and 700 µg/ml G-418 (clonal selection). The assay was
performed in 96-well plates using 5 x 104
cells/100 µl·well in medium containing only 0.5% FCS. Cells were
allowed to adhere overnight, then 100 µl hormones diluted in FCS-free
medium were added to each well. After 24 h of stimulation, cells
were lysed (50 µl), then luciferase activity was counted for 10 sec
in 15 µl cell lysate. The absolute values of luciferase activity were
found to vary depending on the number of cell passages; however, this
did not significantly affect the fold induction of luciferase activity
(calculated as the ratio between the relative light units of stimulated
vs. nonstimulated cells). Maximal fold induction (routinely
15- to 20-fold) was obtained in the range of 1–10 µg/ml WT hPRL.
To avoid interassay variations, all analogs to be compared were
systematically tested within each experiment, and data obtained in one
experiment representative of three experiments performed in duplicate
are shown.
Cell cycle analyses. T-47D cells were routinely cultured in DMEM/Ham’s F-12 (1:1) medium containing 10% FCS, 2 mM glutamine, penicillin (50 U/ml), and streptomycin (50 µg/ml). Cells were aliquoted in six-well plates (0.5–1 x 106 cells/well) and serum-deprived in 0.5% charcoal-stripped serum for 24 h before addition of hormones. After 24-h stimulation, culture medium was collected to recover floating cells by centrifugation, and attached cells were harvested by brief trypsinization. Cells were pooled and permeabilized using 40 µl DNA-Prep Reagent (Coulter Corp., Miami, FL) before 30-min incubation at 37 C in 0.5 ml DNA intercalculator propidium iodide (DNA-Prep stain propidium iodide). Cell cycle distribution was analyzed by flow cytometry on a FACScan using CellQuest software (Becton Dickinson and Co., Mountain View, CA) and manual gating.
Immunoprecipitation and Western blots. Nb2 cells were starved overnight in FCS-free medium before hormonal stimulation. The next day, cells were pelleted and aliquoted in FCS-free medium at 8 x 107 cells/ml (1 ml/tube) and allowed to equilibrate/recover for 30 min at 37 C before PRL stimulation. Cells were stimulated (5 min at 37 C) using various concentrations of WT hPRL or hPRL analogs, as indicated in the figures. At the end of the stimulation, cells were washed twice with ice-cold stopping buffer as previously described (27), and cell pellets were kept frozen until used. Cells were solubilized in 1 ml lysis buffer (30-min rotation at 4 C) (27). Lysates were centrifuged for 10 min at 13,000 x g, then supernatants were quantified for their protein content by Bradford assay and used for immunoprecipitation. A similar procedure was used for Western blot experiments using T-47D cells, except that cells were stimulated for 30 min by the various hormones.
For immunoprecipitation studies (Nb2 cells), 3 mg total lysate were incubated with polyclonal anti-JAK2 (2.5 µl/ml; Upstate Biotechnology, Inc., Schiltigheim, France) or polyclonal anti-STAT5 (C-17, Santa Cruz Biotechnology, Inc., Santa Cruz, CA), used at 5 µl/ml. After overnight rotation at 4 C, immune complexes were captured using 20 µl protein A-Sepharose slurry (Amersham Pharmacia Biotech) for 1 additional h of rotation at 4 C. Protein A complexes were precipitated by centrifugation, and pellets were washed three times in lysis buffer and boiled in 15 µl reducing SDS sample buffer for 5 min at 95 C. Finally, immunoprecipitated samples were analyzed using 7.5% SDS-PAGE. Analysis of MAPK activation was performed on total lysates of T-47D cells using 50–100 µg protein/lane on 10% SDS-PAGE. Electrophoretic transfer onto nitrocellulose membranes (Bio-Rad Laboratories, Inc.) was performed as previously described (27). Membranes were blocked with 5% skimmed milk in Tris-buffered saline-Tween 20 (TBST) for 2 h at room temperature. After washing in TBST, they were incubated overnight (4 C) in 3% BSA/TBST containing 4G10 antiphosphotyrosine antibody (Upstate Biotechnology, Inc.; 1:10,000 dilution) or anti-threonine 202/tyrosine 204-phosphorylated MAPK 1 and 2 (also referred to as antiactive ERK1/2; dilution 1:1,000; New England Biolabs, Inc.). Membranes were again washed in TBST and incubated for 1 h at room temperature with a 1:4,000 dilution of horseradish peroxidase-conjugated antimouse antibody (Amersham Pharmacia Biotech). After washing, immunoblots were revealed by 1-min ECL reaction (Renaissance chemiluminescence kit, NEN Life Science Products, Boston, MA), followed by autoradiography (various exposure times). When required, the membranes were dehybridized as previously described (27). After extensive washing and reblocking, membranes were reprobed using anti-JAK2, anti-STAT5, or anti-MAPK polyclonal antibodies (using antirabbit horseradish peroxidase as secondary antibody). Densitometric analysis of autoradiographies was performed using the image analysis software Scion Image (Scion Corp., Frederick, MD).
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Results |
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represents the
chromatogram obtained for S179D-hPRL. As previously described for
wild-type hPRL (32), the refolded protein is eluted in two
major peaks that were analyzed using reducing and nonreducing SDS-PAGE
(see insets in Fig. 1A). The first peak corresponds to the
void volume of the molecular sieve [elution volume
(Vel), 118 ml] and contains both noncovalent and
disulfide-bonded aggregates of S179D-hPRL and most of the contaminants,
whereas the second corresponds to the monomeric form of the protein
according to column calibration with proteins of known mol wt. Its
Vel (198 ml) is identical to that of monomeric WT
hPRL (23 kDa). Between these two peaks, the chromatogram remains higher
than the baseline, reflecting the presence of low mol wt multimers
(dimers, trimers, etc.). This experiment indicates that
refolded, nonpurified S179D-hPRL [as previously reported for WT hPRL
(32)] is heterogeneous and contains multiple forms of the
protein, which justifies a purification step by gel filtration to
isolate monomeric S179D-hPRL.
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Isoelectric focusing. Figure 1B
represents a typical
isoelectrofocusing experiment of purified WT and hPRL analogs in the pH
range of 3–10. As previously described (32), purified
hPRL exhibits two bands, the major one focusing at pI 6.2, and the
minor one at pI 5.9. According to the addition of a positive charge,
the major band of G129R-hPRL focuses at pH 6.4 as reported previously
(31). Despite the addition of a negative charge,
S179D-hPRL displays the same profile as WT hPRL, with a major
band at pI 6.2 and a minor one at pI 5.9.
Circular dichroism. As point mutations can affect regular
secondary structures and/or global protein folding, we assessed the
-helical content of S179D-hPRL by circular dichroism as previously
reported for other hPRL analogs (31, 33, 40). The profile
obtained for S179D-hPRL was almost identical to that obtained with WT
hPRL (Fig. 1C), with the two minima at 208 and 222 nm and the maximum
at 195 nm typically observed for all -helix proteins, suggesting no
strong (detectable) alteration of the overall content in secondary
structures. The calculated helicity was 45% for WT hPRL and 42% for
S179D-hPRL.
Biological properties
Binding studies. The affinity of S179D-hPRL for the hPRLR was
estimated by its ability to compete with
[125I]hPRL for binding to the receptor. Typical
competition curves are shown in Fig. 1D. The affinity of WT hPRL, as
calculated by Scatchard analysis, indicated a Kd
of 3.4 ± 1.3 x 10-10 M.
As the competition curves of hPRL and S179D-hPRL are almost parallel in
the linear part of the sigmoids (Fig. 1D), the
IC50 ratio reflects the relative affinity of the
mutant for the hPRLR. Averaged from three independent experiments, the
IC50 values of hPRL and S179D-hPRL were 9.5
± 1.0 and 181.7 ± 28.9 ng/ml, respectively, indicating a 20-fold
reduced affinity of the analog for the homologous receptor. As
previously reported (26), the affinity of G129R-hPRL for
hPRLR is 10-fold lower than that of hPRL, i.e. slightly
higher than that of S179D-hPRL.
Bioactivity studies.
Rat Nb2 cells: proliferation assay.
We first tested the
agonistic properties of S179D-hPRL in the Nb2 assay using the classical
procedure recommended by Gout and colleagues (37, 38), who
isolated the Nb2 cell clone and established the lactogen-induced
proliferation assay. Figure 2A shows cell
proliferation induced by increasing concentrations of purified
(left panel) or nonpurified (right panel) WT
hPRL, S179D-hPRL, and G129R-hPRL. As expected, monomeric hPRL induces
cell proliferation with a maximal effect at about 1 ng/ml, whereas the
curve obtained for S179D-hPRL is shifted by 1 log unit toward the
higher concentrations. As previously reported (26, 31),
the agonistic dose-response curve obtained with G129R-hPRL is shifted
by 2 log units compared with that obtained with WT hPRL (Fig. 2A, left). It is noteworthy that the two mutants are able to
induce a maximal proliferation response after 3 d of stimulation
when added at a sufficient concentration; this was confirmed by cell
cycle studies performed using FACS analysis (not shown). Although
the agonistic parts of the curve obtained for nonpurified WT hPRL and
S179D-hPRL (Fig. 2A, right) were similar to their purified
counterparts, they were more markedly bell-shaped at the higher
concentration tested (10 µg/ml). The down-slope in dose-response
curves for PRL and GH receptors is thought to reflect a phenomenon of
self-antagonism (6, 23). However, as no self-antagonism
was observed at the highest concentration with purified proteins (1000
ng/ml in this study, up to 100 µg/ml in Refs. 26, 31
and 33), it is possible that bell-shaped curves obtained
using nonpurified hormones could result from an inhibitory effect of
contaminants present at high doses of nonpurified preparations (see
Fig. 1A and Discussion). Finally, nonpurified G129R-hPRL
failed to induce proliferation.
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50% of maximal cell growth; see Fig. 2A, left), an
increased dose response of proliferation was obtained from
approximately 1 ng/ml of the analog (Fig. 2B, left), which
appeared to be superimposed on the agonistic curve obtained when
testing S179D-hPRL alone (Fig. 2A, left). The same
observation was made for G129R-hPRL (compare left panels of
Fig. 2, A and B). These experiments confirm the intrinsic agonistic
properties of both hPRL mutants in the Nb2 cell proliferation assay.
When 1 ng/ml WT hPRL was competed with nonpurified S179D-hPRL (Fig. 2B, right), the diminution of cell growth observed in the higher
range of analog concentration was also superimposed on the curve
obtained when testing nonpurified S179D-hPRL alone (see Fig. 2A, right), indicating that this effect is not a true
antagonism, but reflects self-antagonism and/or contaminant effects, as
discussed above. On the other hand, the curve obtained with nonpurified
G129R-hPRL (Fig. 2B, right) may reflect true antagonism, as
this protein fraction is intrinsically inactive (Fig. 2A, right). Finally, we noticed that in all of the antagonism
experiments performed using nonpurified proteins (including WT hPRL), a
small decrease in cell response occurred around 0.1–0.2 ng/ml, which
may reflect a slight antagonism at this very narrow range of low
concentrations, although this was not seen with purified preparations
(see Discussion).
Our results are in total contradiction with Chen’s report
(20), which claimed an antagonistic activity for
S179D-hPRL. These researchers performed their Nb2 proliferation assays
using experimental conditions differing from the classical bioassay
protocol, using a lower cell density. Whereas the standard protocol
recommends a cell density of 105 cells/ml on the
starting day (37), Chen et al. used 2.5 x
104 cells/ml for agonistic assays and still less
(5 x 103 cells/ml) for antagonistic assays.
Therefore, we repeated all of our experiments using Chen’s protocol.
With respect to agonism, we obtained dose-response curves qualitatively
similar to those presented in Fig. 2A, except that the changes
in OD450 values of the colorimetric assay
(WST-1 reagent) were lower due to lower cell density (not shown). With
respect to antagonistic assays, although using 1000 cells/well (200
µl) led to very small changes in OD450 values
(<0.1 absorbance unit) that were not very reliable in our hands, we
never observed any evidence for S179D-hPRL antagonism (not shown).
In summary, our results clearly establish S179D-hPRL as a weak agonist, and not an antagonist, in the Nb2 cell proliferation assay regardless of the status of protein purification or cell density.
Activation of the JAK/STAT pathway. Based on the observation that S179D-hPRL activates STAT5 without significantly activating the tyrosine kinase JAK2, Walker’s (28) group proposed that this analog may exert its antagonistic properties by triggering alternate intracellular signals. As in our hands S179D-hPRL failed to display any antagonistic properties, but clearly acts as an agonist, it was important to confirm its effect on the JAK2/STAT5 pathway in Nb2 cells.
Figure 3 shows dose-dependent activation
of JAK2 (A) and STAT5 (B) in Nb2 cells using purified (left
panels) or nonpurified (right panels) hormone
preparations. Regardless of the purification status, S179D-hPRL is able
to activate JAK2 in a manner similar to WT hPRL, which contrasts to the
report of Coss et al. (28). Even at 5 ng/ml, a
faint band corresponding to phosphorylated JAK2 is already visible,
indicating that detection of JAK2 activation does not require
stimulation with supraphysiological concentrations of the analog. In
contrast, G129R-hPRL failed to induce strong activation of JAK2, even
at a concentration of 500 ng/ml (7-fold less phosphorylated JAK2
compared with WT hPRL stimulation; Fig. 3A) or higher (not shown). This
clearly indicates that the threshold of activation of signaling
pathways does not correlate with that of mitogenic activity (compare
Figs. 2 and 3). With respect to STAT5, the same observations were
globally drawn (Fig. 5B), as both hPRL and S179D-hPRL (purified)
achieved a similar level of STAT5 activation. Again, the level of STAT5
activation was markedly lower using G129R-hPRL, as it was when using
nonpurified S179D-hPRL.
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). The only difference between WT
hPRL and S179D-hPRL that we repeatedly observed in these studies was a
lower phosphorylation level of the pool of JAK2 coimmunoprecipitating
with STAT5 (arrowhead in Fig. 3B). This observation was not
analyzed further because it falls out of the scope of this study.
Another finding reported by Coss et al. (28)
was the ability of S179D-hPRL to competitively inhibit hPRL-induced
JAK2 phosphorylation, correlating to some extent with the antagonism
observed in the Nb2 proliferation assay by these researchers
(20). Using similar approaches, we investigated the
antagonistic activity of our S179D-hPRL preparations (purified or not)
on hPRL-induced activation of JAK2 (Fig. 4), using G129R-hPRL as a control.
S179D-hPRL failed to inhibit the effect of 500 ng/ml hPRL on JAK2
phosphorylation even when added in a 50-fold molar excess, and this was
true whether purified or nonpurified analog was used. In contrast, a
50-fold excess of G129R-hPRL clearly competed WT hPRL for JAK2
activation, as the phosphorylation of the kinase was strongly impaired
(by 10-fold) in the presence of the antagonist (Fig. 4, right). The possibility that this inhibition may result from
a toxic effect was ruled out, because no cell death was observed over a
3-d proliferation assay using even higher concentrations of this analog
(31). It is noteworthy that the antagonistic effect
observed on JAK2 was not directly reflected by the level of STAT5
phosphorylation (not shown), which is not fully understood (see
Discussion).
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Transcriptional activity (HL5 clone). We have previously
established a hPRLR-mediated transcriptional bioassay (so-called HL5
clone; see Materials and Methods) that enables us to
unambiguously identify lactogens displaying antagonistic properties
[e.g. mutants of hPRL (25, 26) or hGH
(39)]. In this assay G129R-hPRL fails to act as an
agonist even when tested at extremely high concentrations (120 µg/ml;
Fig. 5A). However, as it binds to the
receptor it acts as an antagonist (Fig. 5B) (26). In
contrast, S179D-hPRL again behaved as a weak agonist, as its curve was
slightly displaced toward high concentrations compared with WT hPRL
(Fig. 4A). Averaged from three experiments, the
ED50 of WT hPRL was 2.3 ± 0.3 µg/ml,
whereas a 4-fold higher concentration of the analog (9.7 ± 0.5
µg/ml) was required to achieve a similar luciferase response.
Surprisingly, the maximal activity obtained at the highest
concentration tested (120 µg/ml) was systematically higher for
purified S179D-hPRL than for WT hPRL (respectively, 23.6 ± 3.0-
and 17.8 ± 1.8-fold inductions; n = 6; Fig. 5A, left). In agreement with its (super) agonistic activity, no
evidence of S179D-hPRL antagonism was observed, as highlighted by the
dose-dependent increase in luciferase activity when the analog and WT
hPRL were tested together (Fig. 5B). These experiments were repeated
using nonpurified material, and similar observations were made,
i.e. that S179D-hPRL activated the hPRLR, whereas G129R-hPRL
did not (not shown).
Proliferation of human mammary tumor cells (T-47D). We
recently reported that G129R-hPRL was able to competitively inhibit
hPRL-induced proliferation of T-47D cells, and this was demonstrated by
monitoring both cell density (WST-1) and cell cycle status in
time-course experiments (27). The same procedure was
repeated here for S179D-hPRL. As expected from the data presented
above, this analog stimulated T-47D cell division, as reflected by the
FACS profile monitored after 24 h of treatment. Due to its lower
affinity for the hPRLR, however, a similar level of cell division
required higher amounts of S179D than WT hPRL (Fig. 6 and data not shown). In agreement with
our previous report, G129R-hPRL (even at high concentrations) failed to
activate T-47D cells. In competition experiments the latter efficiently
competed with hPRL for stimulating entry of T-47D cells into the
division cycle, whereas S179D-hPRL failed to exert any antagonistic
effect.
|
)
(27). In antagonistic experiments, the level of MAPK
activation induced by 100 ng/ml hPRL (leading to mid scale activation
of MAPK; see Fig. 7A) was competed by G129R-hPRL in a dose-dependent
manner. In contrast, S179D-hPRL had an additive effect, leading to
maximal levels of MAPK phosphorylation. These observations clearly
indicate that S179D-hPRL exerts agonistic, but absolutely no
antagonistic, activity on the human PRLR (Fig. 7B).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Upon refolding of inclusion bodies, about 50% of hPRL molecules assemble in high MM aggregates containing both covalent and noncovalent multimers. As PRL polymers have been long known to display reduced biological activity (for a review, see Ref. 41), we routinely eliminate these multimers from refolded hPRL through a single gel filtration step (25, 26, 31, 32, 33, 42). As illustrated by SDS-PAGE analysis, this step also eliminates most contaminants. In contrast, Walker et al. performed all of their studies using refolded, nonchromatographically purified S179D-hPRL. These researchers justified omitting the purification step by claiming that limiting the time of protein overexpression (IPTG) to 2 h, instead of 4 h as recommended in the initial protocol (32), reduces the amount of high MM contaminants pelleting within inclusion bodies (20), and their protein preparations did not contain oligomeric forms based on nonreducing SDS-PAGE analysis (data not shown in Ref. 20). We disagree with the latter argument, as noncovalent multimers cannot be detected by denaturing electrophoresis. Moreover, as we obtained indistinguishable chromatograms whether IPTG induction was performed for 2 h (not shown) or 4 h, we anticipate that the nonpurified hormone preparations used by Chen and colleagues are heterogeneous and contain a significantly amount of high MM proteins. Unfortunately, due to the absence of a gel filtration profile in their report, the true monomer/multimer ratio of their preparations cannot be assessed. Of more concern is that our nonpurified fraction probably contains bacterial contaminants normally eluted in the void volume of the molecular sieve, which may interfere with bioassays. The pronounced bell-shaped curves observed for nonpurified preparations compared with their purified counterparts may actually reflect such a PRL-independent effect resulting from bacterial contaminants presumably present in significant amount at high doses of hPRL analogs. Our policy is to always use purified proteins for characterizing hPRL analogs. Therefore, we will not go into detailed analysis of data obtained using our nonpurified material, because interpretation of such results could be misleading. Their use was solely to assess that the purification parameter alone did not support the contradiction between Walker’s and our observations. Our data conclusively show that nonpurified S179D-hPRL acts as an agonist in the Nb2 proliferation assay. These observations were further confirmed, first using the luciferase bioassay developed in our laboratory and mediated by the homologous (human) PRLR, and second by monitoring T-47D cell proliferation or activation of the MAPK cascade known to be triggered by PRL in this human mammary tumor cell line. Depending on bioassay sensitivity, S179D-hPRL was clearly able to activate the hPRLR with an efficiency decreased by approximately 1 log unit, in perfect agreement with its 20-fold reduced binding affinity for this receptor. In summary, the discrepancy between both reports cannot result from different intrinsic properties of the protein preparations used.
For 20 yr, monitoring Nb2 cell proliferation has been established as the reference bioassay for quantifying lactogen activity (37, 43). Although the classical experimental procedures recommend using 105 cells/ml on the starting day of the assay, Walker and colleagues used 4 times less cell density in their bioassays, which we assumed may have influenced some characteristics of cell responses to hormone stimulation. Supporting this hypothesis, Chen et al. (20) reported unusual dose-response curves, with maximal response peaking at 200–400 pg/ml, whereas normally maximal cell growth is seen at 1–2 ng/ml hPRL (37). Still more confusing, they observed hPRL self-antagonism (reflected by bell-shaped curves) at concentrations as low as 1.6 ng/ml (20), whereas we (31) and many others (44, 45, 46, 47) failed to detect any self-antagonism of lactogens on Nb2 cells at much more elevated concentrations (up to 1 µg/ml). To clarify this issue, we repeated all of our experiments following Chen’s protocol (2.5 x 104 cells/ml), and our results appeared qualitatively similar (not shown), i.e. that S179D-hPRL (purified or not) acted as a weak agonist on Nb2 cell proliferation.
As a first conclusion regarding agonistic assays, all of our data support that the agonistic properties of S179D-hPRL are real and were missed in Walker’s reports. The most probable reason for this misinterpretation is that their bioassays were performed using an inappropriate concentration range of ligand. In fact, the growth-promoting effect of S179D-hPRL in the Nb2 assay occurs between 1–100 ng/ml, whereas the concentrations tested by Chen and colleagues did not exceed 1.6 ng/ml. This hypothesis was definitely confirmed when a S179D-hPRL preparation provided by Walker was shown in our hands to exhibit agonistic properties very similar to those of our preparation (not shown) in both Nb2 and HL5 assays.
As discussed previously (25), Nb2 cell proliferation is an extremely sensitive bioassay and therefore is probably not the most appropriate bioassay for highlighting the antagonistic properties of hPRL analogs. For example, although G129R-hPRL (25, 26, 27) and G120R-hGH (or G120K-hGH) (24, 39, 48) have been characterized as potent antagonists toward the PRLR in various cell-based assays, both are weak agonists and fail to antagonize WT lactogens in the Nb2 cell proliferation assay (31, 39, 44). This can be explained by the fact that maximal Nb2 cell proliferation is achieved at very low receptor occupancy (49), as highlighted by immunoblot analyses where maximal activation of signal transducers (e.g. JAK2 and STAT5) requires concentrations as high as 500 ng/ml, when 1 ng/ml hPRL is sufficient to promote maximal cell proliferation. Hence, PRL/GH analogs whose ability to dimerize the receptor is strongly, but not totally, abolished maintain the ability to induce receptor dimer formation to a minimal level, sufficient to promote Nb2 cell division (50). In view of this experimental background, the highly potent antagonistic properties of S179D-hPRL in the Nb2 assay reported in Chen’s study appeared intriguing (20). They were still more questionable in view of the agonistic properties exhibited by S179D-hPRL in our hands. Expectedly, we failed to observe any evidence for antagonism when this analog was tested in competition experiments in the Nb2 assay. To circumvent the paradoxical response of Nb2 cells to hPRL analogs otherwise acting as potent antagonists, we have recently developed the PRLR-LHRE transcriptional assay that previously allowed unambiguous identification of PRL/GH mutants exhibiting antagonistic properties (25, 26, 39). As the transcriptional response, in contrast to cell division, is not (or minimally) affected by intracellular limiting factors, one can consider that the luciferase dose-response curve parallels that of receptor dimer formation (25, 50). In this bioassay, S179D-hPRL clearly lacks any antagonistic activity, whereas G129R-hPRL, the most potent hPRL antagonist currently available, does antagonize WT hPRL. These observations were further strengthened using human mammary T-47D cells. We (27) and others (48) have previously shown that PRL antagonists inhibit lactogen-induced proliferation of mammary tumor cells as well as the major PRLR-triggered signaling cascades, including JAK2; STAT1, -3, and -5; and MAPKs (27). In this study we monitored the inhibition of MAPK by hPRL analogs. Clearly, whereas G129R-hPRL leads to extinction of the phosphorylation signal induced by the intermediate dose of hPRL, S179D-hPRL still increases MAPK activation. Therefore, we must conclude that S179D-hPRL is completely devoid of any detectable antagonistic activity. To understand why contradictory results were obtained, we repeated the experiments involving Nb2 cells using the protocol described in Chen’s study (20). In our hands, using a cell density as low as 5 x 103 cells/ml appeared unreliable, as the OD450 values were so low that the variation in absorbance (change in OD450, <0.1 U) was not significant. We repeatedly observed a small decrease in Nb2 cell response in a very narrow range of low concentrations using nonpurified hPRLs (including WT hPRL) in competition experiments. Although we cannot propose any molecular mechanism to support this observation, such a phenomenon limited in amplitude and in concentration range may be reminiscent of what Walker and colleagues interpreted as reflecting S179D-hPRL antagonism.
In a second report (28) the same group suggested that
S179D-hPRL may activate alternative intracellular signaling cascades,
although such pathways were not identified. In a more general fashion
their data argued for a dissociation between JAK2 and STAT5 activation,
in total contradiction with the conventional model of JAK-induced
phosphorylation of STATs (51). Again, we absolutely failed
to confirm these observations, as S179D-hPRL, purified or not, appeared
able to induce JAK2 phosphorylation in a fashion similar to WT hPRL in
Nb2 cells. A tentative explanation for the discrepancy between the
results of Walker’s group and ours might be that the level of JAK2
phosphorylation was, in general, low in their hands, but detectable
(Fig. 3A in Ref. 28), which may have led the researchers
to suggest the involvement of another kinase in STAT5 activation.
Accordingly, we obtained stronger signals using the anti-JAK2 antibody
from Upstate Biotechnology, Inc., compared with that from
Santa Cruz Biotechnology, Inc., used in their study (data
not shown), which, however, does not support the discrepancy, as the
antibody from Santa Cruz Biotechnology, Inc.,
successfully detected JAK2 phosphorylation induced by WT hPRL
(28). Moreover, we observed that G129R-hPRL induced only a
limited level of JAK2 phosphorylation, demonstrating that our
experimental procedures were sensitive enough to identify analogs
exhibiting lower ability to activate this signaling cascade. According
to these observations regarding agonistic effect, we failed to confirm
the antagonistic activity of S179D-hPRL on hPRL-induced JAK2 activation
previously reported by these researchers, whereas the antagonism of
G129R-hPRL could be detected, in agreement with its inhibitory effect
on PRLR signaling pathways recently demonstrated in breast cancer cells
(this study and Ref. 27). The only difference repeatedly
observed between S179D-hPRL and WT hPRL is a decrease in the pool of
phosphorylated JAK2 coprecipitating with STAT5, which may reflect a
difference in the stoichiometry and/or conformation of PRLR/JAK2/STAT5
complexes depending on the ligand (see below). Obviously, the
importance of this observation will require further investigation,
which, however, is out of the scope of the present study.
All of the results presented in this study clearly establish S179D-hPRL as an agonist (although weaker than hPRL), not as an antagonist, of the PRL receptor. However, our report raises many questions, the first of which is how this hPRL analog should be categorized. Several arguments suggest that S179D-hPRL is a binding site 1 rather than a binding site 2 analog. Based on the three-dimensional model that we established (52), serine 179 is predicted to belong to helix 4, the region of binding site 1 containing the highest number of binding determinants (6). Moreover, the dose-response curves obtained for S179D-hPRL in the various bioassays investigated here are displaced to the right compared with WT hPRL and achieve a maximal response (or supramaximal, see below), two features that fit with alteration of binding site 1 (6, 33, 42, 50). Although S179D-hPRL can be considered as a site 1 analog, this does not necessarily mean that serine 179 is intrinsically involved in an interaction with the PRLR. One of the most powerful methods for identifying binding determinants of PRLs or GHs has proved to be alanine scanning (for review, see Ref. 6). Although Chen et al. (20) reported that alanine replacement of serine 179 abolishes hPRL activity, which would argue that this residue is functionally relevant, we cannot exclude that the putative agonistic activity of the S179A-hPRL analog could have been missed, as it was for S179D-hPRL (see above). In contrast to all binding determinants identified to date (6), serine 179 is predicted to point toward the hydrophobic core of the protein (52), which presumably precludes this residue from directly interacting with the receptor. This structural hypothesis is corroborated by isoelectric focusing experiments showing that the pI of S179D-hPRL is identical to that of WT hPRL despite the addition of a negative charge (Asp for Ser). We expect that Asp179 (and Ser in WT hPRL) is buried and that its putative charge does not influence electrophoretic mobility. If true, this would suggest that the reduced agonism of S179D-hPRL would not result from the mutation of Ser179 per se, but, rather, from the effect this mutation has on overall protein structure. The estimation of S179D-hPRL apparent MM using analytical gel filtration did not reveal any detectable change with respect to WT hPRL, but this approach is obviously not very sensitive to small structural variations. Circular dichroism previously enabled us to detect structural disturbance of hPRL analogs (e.g. C58A or S26W) (31, 33). Using this approach, we failed to detect any significant alteration of secondary structure content of S179D-hPRL, as its spectrum and calculated helicity were not significantly different from those of WT hPRL (this study and Refs. 31 and 33). In agreement with Chen and colleagues, who used RIA recognition to monitor conformational disturbances, we thus conclude that the S179D substitution does not dramatically influence the overall hPRL structure.
As Ser179 is presumably buried and, in addition, mutation of this residue leads to reduction of affinity and of agonistic activity without any detectable structural alteration, we hypothesize that aspartate substitution results in local disturbance of regions involved in binding site 1, most probably of helix 4. Recent investigation of Walker’s group indicates that Ser179 can be phosphorylated in vitro in the human hormone (Lorenson, M. Y., and A. Walker, personal communication), as already demonstrated for the homologous Ser177 in rPRL (18), which, however, does not mean that these residues actually are phosphorylated in the pituitary in vivo. Elucidating the mechanism by which an amino acid predicted to be buried inside the protein becomes accessible to a kinase remains open to investigation. Whether S179D-hPRL actually exhibits the same properties as a putative S179-phosphorylated hPRL, thereby justifying being referred to as a molecular mimic of phosphorylated hPRL remains an open question (20, 53).
The final and probably most important questions relate to the intrinsic
properties that S179D-hPRL exerts in vivo when injected to
animals. Within the last year, several reports on the effect of
S179D-hPRL in various rodent models have been presented.
Nonexhaustively, S179D-hPRL was reported to reduce the tumor incidence
of cancer prostate cells injected into nude mice (54) and
in rats to alter maternal behavior in nulliparous female
(55) and to affect T cell survival (56),
which is in good agreement with the antagonistic properties claimed by
Walker and colleagues from their in vitro studies (20, 28), but obviously does not agree with our conclusions. In
contrast, S179D-hPRL was also shown to promote lobuloalveolar
differentiation and casein expression during rat pregnancy
(57). Moreover, a very recent publication by the same
group described S179D-hPRL as an even more potent agonist than WT hPRL
on the inhibition of osteoblastic alkaline phosphatase activity and
bone development in rats (58), which fits with the
agonistic activity demonstrated for this analog in the present work.
Although one of our concerns about the relevance of these preliminary
in vivo studies is whether nonpurified proteins obtained
from bacteria are a suitable material to be injected into animals,
especially for investigations of immunological parameters, the
paradoxical properties of S179D-hPRL both in vitro and
in vivo obviously argue that this hPRL mutant should be
viewed with much caution and certainly not as a full antagonist as
suggested previously (20, 55, 56). Unexpectedly, our
transcriptional bioassay revealed that although S179D-hPRL behaves as a
slightly weaker agonist than hPRL (dose-response curve displaced to the
right), it induces a luciferase response of higher amplitude (125%)
compared with WT hPRL, thereby displaying superagonistic properties
in vitro. As this phenomenon was also observed using the
hormone preparation provided by Walker (not shown), it may correlate
with the superagonistic activity reported for S179D-hPRL on the
inhibition of bone formation (58), although this reported
effect would be opposed to the reduced bone development we have
reported in PRL receptor knockout mice (59). This suggests
that once the functional hormone-receptor complex is formed (which
requires higher hormone concentration due to reduced affinity),
S179D-hPRL might be able to trigger signaling cascades more efficiently
than the natural ligand. Whether this property is related to prolonged
signal transduction, a different
kon/koff affinity constant,
conformational changes in the PRLR, or some other molecular feature
remains to be explored.
In conclusion, the present report clearly establishes S179D-hPRL as a PRLR agonist and suggests that the antagonistic properties claimed for this analog were deduced from inappropriate bioassays. Although it is assumed that phosphorylated PRL plays a role in autoregulation of PRL secretion from the pituitary or is an intracellular component acting as a regulator of trafficking and/or storage of PRL molecules, the physiological relevance of this PRL variant remains poorly understood (12). Based on injection of S179D-hPRL to pregnant rats, it was recently proposed that the ratio of nonphosphorylated vs. phosphorylated hPRL, rather the latter per se, may be physiologically relevant. We strongly feel that extrapolation of findings with S179D-hPRL to the potential physiological relevance of phosphorylated hPRL in vivo remains to be explored and should await the definite demonstration that the S179D mutation actually confers properties similar to those specifically exhibited by phosphorylated PRL.
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Acknowledgments |
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Footnotes |
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Abbreviations: hPRL, Human PRL; IPTG, isopropylthiogalactoside; JAK, Janus kinase; LHRE, lactogenic hormone response element; MM, molecular mass; PRLR, PRL receptor; rPRL, rat PRL; STAT, signal transducer and activator of transcription; TBST, Tris-buffered saline-Tween 20; Vel, elution volume; WT, wild type.
Received December 6, 2000.
Accepted for publication May 9, 2001.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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) is expressed in degree x
cm2 x dM-1. Both spectra
exhibit the profile typical of all 
) or
S179D-hPRL analog (•). Each point is the average of
duplicate measurements calculated from three independent experiments
(error bars represent the SD). Based on the
IC50, S179D-hPRL displays an affinity 20-fold lower than
that of WT hPRL.
) were performed as indicated in
Materials and Methods, using 105 cells/ml on
the starting day. Cell proliferation was assessed using WST-1
colorimetric reagent. The data presented are representative of at least
three independent experiments that were performed in triplicate or
quadruplicate. Error bars represent the SD.
A, Agonism. Cell proliferation induced by WT hPRL occurs in the range
of 0.1–1 ng/ml. Proliferation curves observed in the presence of
purified S179D-hPRL or G129R-hPRL are displaced to the high
concentration by 1 and 2 log units, respectively, and reach maximal
proliferation (same plateau as WT hPRL). Nonpurified WT hPRL and
S179D-hPRL lead to bell-shaped curves, whereas nonpurified G129R-hPRL
fails to induce cell growth. B, Antagonism. Purified hPRL analogs
failed to antagonize cell division induced by 200 pg/ml WT hPRL. The
decrease in cell growth in the presence of high concentrations of
nonpurified analogs is discussed in the text.
