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Endocrinology Vol. 148, No. 3 1330-1339
Copyright © 2007 by The Endocrine Society

C-Terminal Fragments of the Gastrin-Releasing Peptide Precursor Stimulate Cell Proliferation via a Novel Receptor

Oneel Patel, Chelsea Dumesny, Arthur Shulkes and Graham S. Baldwin

Department of Surgery, University of Melbourne, Austin Health, Heidelberg, Victoria 3084, Australia

Address all correspondence and requests for reprints to: Graham S. Baldwin, Department of Surgery, Austin Health, Studley Road, Heidelberg, Victoria 3084, Australia. E-mail: grahamsb{at}unimelb.edu.au.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
There are many precedents for the production from a single precursor of multiple peptides, with independent receptors and different bioactivities. Gastrin-releasing peptide (GRP) is initially synthesized as amino acids 1–27 of a 125-residue precursor, proGRP, and is subsequently cleaved and amidated to form GRP18–27. We investigated the hypothesis that C-terminal proGRP peptides are also biologically active. Human proGRP18–125 was expressed in Escherichia coli as a glutathione S-transferase fusion protein. Recombinant proGRP18–125 stimulated proliferation and migration of the human colorectal carcinoma cell line DLD-1. The observations that an antagonist selective for the GRP receptor did not inhibit activity in either proliferation or migration assays and that the recombinant peptide did not bind to either the GRP receptor or orphan receptor BRS-3 indicated that neither activity was mediated by the known GRP receptors. Recombinant human proGRP31–125 and proGRP42–98 were also prepared and shown to stimulate proliferation of DLD-1 cells and the human prostate carcinoma cell line DU145. The synthetic peptides proGRP47–68 and [Tyr79]proGRP80–97 stimulated inositol phosphate production, MAPK kinase activity, and proliferation and migration of DLD-1 cells. Binding sites for both radioiodinated synthetic peptides were demonstrated on DLD-1 cells. Each peptide was able to compete with the other for binding, and a GRP receptor antagonist did not inhibit binding of either peptide. We conclude that peptides derived from the C terminus of proGRP are biologically active and that their activity is mediated by a receptor distinct from the two known GRP receptors.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GASTRIN-RELEASING PEPTIDE (GRP) was initially discovered in the stomach and named for its potent stimulation of gastrin release (1). However, the widespread distribution of GRP, with significant amounts present in the central nervous system and throughout the entire gastrointestinal tract, suggests more general actions. GRP is now known to perform many biological functions that are independent of its stimulation of gastrin release (2). In particular GRP has been recognized as the prototypical autocrine growth factor, based on the detection of GRP and its cognate receptor in small cell lung carcinoma (SCLC) and on the antiproliferative effect of GRP antibodies (3). GRP is also a potent mitogen for several other types of carcinomas including colorectal, pancreas, prostate, and breast (4, 5).

All forms of GRP are derived from a 125-amino acid precursor, proGRP (Fig. 1Go). The generation of amidated forms of GRP from the precursor proGRP begins with cleavage after Lys29Lys30 by a prohormone convertase. Carboxypeptidase E then removes the lysine residues to generate glycine-extended GRPs (GRPglys), which may be converted to C-terminally amidated GRP1–27 by peptidyl {alpha}-amidating monooxygenase. Further endoproteolytic cleavage after Arg17 releases the amidated decapeptide GRP18–27. Both GRP1–27 and GRP18–27 have C-terminal sequences identical with the frog peptide bombesin (1). Evidence is also accumulating that nonamidated forms of GRP18–27 are biologically active and are present in normal and tumor tissues. Oiry et al. (6) reported that a synthetic glycine-extended bombesin is as biologically active as amidated GRP18–27 in fibroblasts and a pancreatic cell line, and we have shown that GRPgly stimulates proliferation of both the colorectal cancer cell line DLD-1 (7) and the colorectal mucosa in an animal model (8). We also expressed proGRP1–125 as a His-tagged fusion protein in Escherichia coli and shown that the HPLC-purified prohormone stimulated MAPK activity and proliferation in DLD-1 cells (9).


Figure 1
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  		FIG. 1. Processing of proGRP. PreproGRP (148 amino acids) is converted to proGRP (125 amino acids) by removal of the signal peptide. The sequential actions of a prohormone convertase that cleaves on the C-terminal side of the sole pair of lysine residues, and a carboxypeptidase B-like enzyme that removes the lysines, convert proGRP to glycine-extended proGRP1–27. The C terminus of proGRP1–27 is then amidated by peptidyl {alpha}-amidating monooxygenase. The C-terminal fragments of proGRP (proGRP18–125, proGRP31–125, and proGRP42–98) were all expressed in E. coli.

 
The three known receptors for peptides of the GRP family, the GRP receptor (GRP-R), neuromedin B (NMB) receptor, and bombesin receptor subtype 3(BRS-3), are members of the G protein-coupled superfamily of receptors with seven-transmembrane domains. GRP-R has a high affinity for GRP18–27, and a widespread distribution in the central nervous system and gastrointestinal tract. The NMB-R has a 2500-fold higher affinity for NMB than GRP (5) and is expressed in the brain and esophagus (10). In contrast, no high-affinity, naturally occurring ligand for BRS-3 has been identified. BRS-3 has low affinity for GRP18–27 and is found in the brain, spinal cord, and uterus (11). The affinity of GRP-R for GRP18–27gly is approximately 20-fold lower than for amidated GRP18–27 (7). Mice deficient in GRP-R have increased body weight (12), a complete attenuation of colorectal villous growth between d N1 and N12 (13) and more persistent long-term fear memory (14) than wild-type controls. The observation that mice lacking functional BRS-3 develop a mild obesity associated with hypertension and impairment of glucose metabolism suggests that BRS-3 is involved in the endocrine control of energy balance and adiposity (15). Several high-affinity peptide antagonists have been developed for GRP-R (16), and these have been effective in a number of animal models of cancer (5).

Although it has been assumed that bioactive GRP peptides are derived exclusively from residues 1–27 of proGRP, there are many precedents for the production from a single precursor of multiple peptides, with independent receptors and different bioactivities. For example, glucagon, glucagon-like peptide 1, and glucagon-like peptide 2 are all cleaved from the 160-amino acid precursor proglucagon (17). Each glucagon-like peptide has its own distinct receptor, and the three receptors are encoded by separate genes at different chromosomal locations (17). The possibility that bioactive peptides might also be derived from residues 31–125 of the precursor proGRP (Fig. 1Go) has not been systematically investigated, although such C-terminal forms of proGRP are present in normal and tumor tissues. High-molecular-weight forms including proGRP31–98 are present in the tumors and circulation of patients with SCLC (18), and an assay that measures circulating proGRP31–98 has been developed for cancer diagnosis and treatment monitoring (19). Because processing of peptides in tumors is often incomplete and secretion is unregulated (20), the possibility should be borne in mind that nonamidated proGRP-derived peptides may be important in cancer biology.

To test the hypothesis that the C-terminal region of proGRP is biologically active, we expressed proGRP18–125, and various peptides derived from it, in Escherichia coli. The recombinant peptides have been purified, characterized, and tested for biological activity in cell proliferation and migration assays and their ability to bind to the GRP and BRS-3 receptors. Synthetic peptides derived from the C-terminal region of proGRP have also been tested in the same assays.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
The cell lines used were obtained from the following sources: DLD-1, HCT15, HT-29, DU145, and Ishikawa (American Type Culture Collection, Manassas, VA); SW1222 (Ludwig Institute for Cancer Research, Melbourne, Australia). DLD-1 and HEK 293 cells were maintained in DMEM with 10% fetal bovine serum (FBS), HCT15, HT29, and SW1222 cells in RPMI 1640 with 10% FBS, and DU145 and Ishikawa cells in MEM with 10% FBS. All cells were grown in 5% CO2 at 37 C. Bombesin and amidated GRP were obtained from Auspep (Melbourne, Australia). GRPgly (purity > 99%) was custom synthesized by Auspep; proGRP47–68 (purity > 93%) and [Tyr79]proGRP80–97 (purity > 95%) were custom synthesized by Mimotopes (Melbourne, Australia) A tyrosine residue was placed at the N terminus of proGRP80–97 to permit radioactive iodination for binding experiments. No such extension was necessary in the case of proGRP47–68, which already contained a tyrosine residue at position 51. Peptide composition and purity were determined by mass spectrometry and HPLC, respectively. The GRP-R receptor antagonist [D-Phe6, Leu-NHet13, des-Met14]-bombesin-6–13 was purchased from Bachem AG (Bubendorf, Switzerland).

Construction of plasmids encoding proGRP
The DNA-encoding proGRP1–125 was amplified by PCR with forward primer GRP35 and reverse primer GRP36 (Table 1Go) and the DNA encoding human preproGRP1–125 cloned into the pSp64 vector (a generous gift from Professor James Battey, National Institute for Deafness and Other Communication Disorders, Bethesda, MD) as template. DNA was amplified using Pfx polymerase (Invitrogen, Groningen, The Netherlands), and the PCR product of 410 bp was ligated into the pGEX-2TH vector.


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  		TABLE 1. Primers used for amplification of DNA encoding C-terminal proGRP fragments

 
The DNA encoding proGRP31–125 was cloned into the vector pGEX-2TH by overlap extension PCR (21). In the first PCR, a forward primer (pGEXupstream, Table 1Go) located in the region encoding GST and the proGRP1 reverse primer were used with the proGRP1–125 clone in pGEX-2TH as template to generate a 144-bp PCR product. A second PCR with the proGRP2 forward primer and the GRP36 reverse primer gave an amplified product of 282 bp. The PCR products from reactions 1 and 2 were combined together in a 2:1 ratio and used as template in a third PCR with the pGEX upstream forward primer and the GRP 36 reverse primer. The final PCR product of 426 bp was ligated into the pGEX-2TH vector.

The DNA encoding proGRP42–98 was amplified using forward primer proGRP3 and reverse primer proGRP4 (Table 1Go) and the proGRP1–125 clone in pGEX-2TH as template. The initial PCR was carried out using Pfx polymerase, but because the product yield was very low subsequent, PCR was carried out using Taq polymerase. The PCR product of approximately 170 bp was ligated into pGEX-2TH vector.

Each of the plasmid DNAs was electroporated into electrocompetent E. coli strain DH5{alpha} for plasmid maintenance and strain BL21 for protein expression. Clones were sequenced using an upstream pGEX-2TH-vector forward primer to confirm that there were no mutations within the proGRP sequence.

Expression, purification, and characterization of proGRP fragments
The GST-proGRP fusion proteins were expressed in E. coli strain BL21 and purified from sarkosyl lysates by chromatography on glutathione-agarose as described previously (22). The fusion protein was then eluted from the beads by treatment with 5 mM glutathione for 1 h at 4 C and digested with thrombin (4 U per 2 mg of fusion protein) for 3 min at 37 C. These conditions were found to minimize the proportion of fusion protein cleavage at the Lys79-Ala80 bond in proGRP. The thrombin digest was loaded on to a C18 µbondapak column (8 x 100 mm; Waters, Rydalmere, Australia), which had previously been equilibrated in 0.05% trifluoroacetic acid, and eluted with the indicated gradient from 0 to 70% acetonitrile in 0.05% trifluoroacetic acid over 60 min at a flow rate of 1 ml/min. Fractions of 1 ml were collected and their protein content analyzed by PAGE. Fractions containing proGRP fragments were dried with a Speed-Vac (Savant, Hicksville, NY). N-terminal sequence and molecular mass were determined as described previously (22) by sequential Edman degradation and electrospray ionization mass spectrometry, respectively.

Cell proliferation assays
Cell proliferation was measured using the 3-[4,5-dimethylthiazol-2-yl]2,5-diphenyltetrazolium bromide (MTT) colorimetric assay, which is based on the reduction of MTT to a blue formazan product by mitochondrial dehydrogenases present in viable cells (23). Briefly, 1 x 104 cells/well were plated into a 96-well plate in DMEM containing 10% FBS. The next day the medium was replaced with serum-free medium and cells were incubated for a further 24 h. On the third day, peptides were diluted in medium containing 0.2% FBS, added to the cells, and incubated for a further 3 d. The total volume in each well was 200 µl. At the end of incubation 15 µl MTT solution (5 mg/ml in PBS) was added to each well and incubated at 37 C for 4 h. The medium was then removed carefully without disturbing the insoluble formazan crystals. Two hundred microliters of 0.04 M HCl in isopropanol were added to each well to solubilize the crystals, and the absorbance was read at 570 nm using a spectrophotometer. Readings from wells that received control medium and no MTT treatment were used as blanks.

Cell proliferation was also measured by [3H]thymidine incorporation. Briefly, 1 x 105 cells/well were plated into a 24-well plate in DMEM with 10% FBS. The next day the cells were synchronized in G0 by starving the cells in serum-free medium for 24 h. On the third day, test peptides were diluted in DMEM containing 1% FBS, added to cells, and further incubated for 17 h. The cells were then pulsed with 0.5 µCi of [3H]thymidine for 4 h and incorporated radioactivity measured as described previously (9).

Wound-healing assay
To study the migratory effect of peptides, wound-healing assays were performed as described previously (24). Briefly, cells were seeded in 12-well plates in DMEM containing 10% FBS and grown until 90% confluent. Cells were then serum starved for 24 h, and a linear wound 1.00 ± 0.02 mm wide was created in the confluent monolayer using a 20-µl pipette tip. Cells were then washed with PBS and treated with or without different peptides diluted in DMEM containing 0.2% FBS. Wounds were photographed at 0, 17, and 24 h, and the wound size was measured at five random sites perpendicular to the wound.

Inositol phosphate production assay
The 5 x 104 cells/well were seeded in DMEM containing 10% FBS in a 24-well plate and allowed to attach overnight. The next day cells were labeled with [3H]myoinositol for 24 h in serum-free medium. The cells were then incubated with or without peptides diluted in assay buffer [20 mM HEPES, 135 mM NaCl, 2 mM CaCl2, 1.2 mM MgSO4, 1 mM EGTA, 10 mM LiCl, 11.1 mM glucose, 0.5% BSA (pH 7.45)] for 1 h at 37 C. The reaction was stopped by adding 1 ml of acidified ethanol [1:2000 (vol/vol) concentrated HCl/ethanol], and the solution was then loaded on to a Dowex AG-1-X8 column (formate form) (Bio-Rad, Hercules, CA). Columns were washed with 4 ml distilled water followed by 4 ml of 40 mM ammonium formate. Inositol phosphates were eluted with 4 ml of 1 M ammonium formate, and radioactivity was measured after addition of 5 ml of scintillation fluid to the eluates.

MAPK activation assay
Cells were grown in 10-cm petri dishes in DMEM containing 10% FBS until 90% confluent. After serum starvation, the cells were stimulated for 5 min with 10 nM proGRP42–98, proGRP47–68, or [Tyr79]proGRP80–97. Cells were lysed and lysate protein (20 µg) separated by electrophoresis and Western blotted with antibodies against either total or phosphorylated p42/p44 MAPK as described previously (9).

Receptor binding assay
Receptor binding assays were performed with BALB3T3 fibroblasts that had been stably transfected with plasmids encoding either GRP-R or BRS-3 (a generous gift from Professor James Battey). Binding of proGRP fragments to GRP-R and BRS-3 was measured by competition with 125I-[Tyr4] bombesin or the 125I-BRS-3 agonist [D-Tyr6, ß-Ala11, Phe13, Nle14]bombesin-6–14, respectively. Briefly 1.5 x 105 cells/well for each cell line were plated in medium containing 10% FBS. Cells were serum starved for 24 h. The next day, cells were washed once in binding buffer [200 mM TrisCl, 100 mM KCl, 2 mM MgCl2, 1 mM DTT, 1 mM benzamidine, 0.1% BSA (pH 7.2)]. Cells were incubated with labeled bombesin or labeled BRS-3 agonist for 45 min at 37 C with or without 1 µM unlabeled bombesin. At the end of the incubation, cells were washed and radioactivity measured as described previously (9).

Binding assays for proGRP fragments
ProGRP47–68 and [Tyr79]proGRP80–97 were individually radioiodinated with 125I (specific activity 3300 cpm/fmol; ICN Biomedicals, Seven Hills, Australia) using the chloramine-T oxidation method. Both radiolabeled peptides were purified by reverse-phase HPLC. Binding of iodinated proGRP fragments to DLD-1 cells was investigated and specificity measured by competition with unlabeled peptides. Briefly, each cell line was plated at 5 x 105 cells/well in medium containing 10% FBS. The next day, cells were washed once with binding buffer. Cells were incubated with labeled proGRP47–68 or [Tyr79]proGRP80–97 (1.5–4.5 x 105 cpm) in 0.5 ml binding buffer for 45 min at 37 C with or without the different peptides (see Fig. 6Go). At the end of the incubation, cells were washed and radioactivity measured as described previously (9).


Figure 6
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  		FIG. 6. Affinity of C-terminal proGRP fragments. Binding of 50 pM 125I-labeled proGRP47–68 (A) or 125I-labeled [Tyr79]proGRP80–97 (B) to 1.5 x 104 DLD-1 human colorectal carcinoma cells was measured in the presence of increasing concentrations of unlabeled proGRP31–125 ({blacktriangleup}, solid line), proGRP42–98 ({triangleup}, dotted line), proGRP47–68 (bullet, solid line), or [Tyr79]proGRP80–97 ({circ}, dotted line), as described in Materials and Methods. Points represent experimental data expressed as a percentage of the radioactivity bound in the absence of any unlabeled peptide (i.e. percent control). Points are mean ± SEM of at least three experiments, each in duplicate. Lines of best fit were obtained with the program Sigmaplot and Kd values are given in Results. For proGRP47–68 (A) a one-site model (maximum binding 100 ± 3%; minimum binding 8 ± 3%; Kd 170 ± 30 nM; R2, 0.98) gave the best fit, whereas for [Tyr79]proGRP80–97 (B), a two-site model [maximum binding 95 ± 6%; minimum binding 14 ± 3%; Kd1 40 ± 50 fM (32%) and Kd2 38 ± 12 nM (68%); R2, 0.97] was superior to a one-site model (maximum binding 74 ± 2%; minimum binding 16 ± 4%; Kd 24 ± 11 nM; R2, 0.91). The minimum binding values of 8 and 14% were then used in the fitting of the data for the other peptides in A and B, respectively.

 
Statistics
Results are expressed as the means ± SEM. Data were analyzed by one-way ANOVA. If there was a statistically significant difference in the data set, individual values were compared with the control by t test with Bonferroni correction for multiple analyses. Differences with P < 0.05 were considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Conservation of proGRP31–125 sequences
Comparison of the sequences of the C-terminal region of proGRP reveals extensive homology among human, cow, sheep, rat, and mouse. Such similarity is often a consequence of the evolutionary pressure maintained by the need for a peptide to bind to its cognate receptor but alternatively might indicate an important role in proGRP secretion. To test the hypothesis that the C-terminal region of proGRP is biologically active, we expressed proGRP, and various peptides derived from it, in E. coli.

Expression and characterization of C-terminal proGRP fragments
proGRP18–125, proGRP31–125, and proGRP42–98 were independently expressed as fusion proteins with glutathione S-transferase in E. coli strain BL21 as described in Materials and Methods. The fusion proteins were isolated by binding to and elution from glutathione-agarose beads and cleaved with thrombin. The thrombin digest containing proGRP31–125 was separated by reverse-phase HPLC (Fig. 2AGo), and subsequent analysis by PAGE (Fig. 2BGo) revealed that the peak at 25 min marked with an asterisk contained proGRP31–125. Because thrombin cleaves on the C-terminal side of Pro-Lys or Pro-Arg sequences, the peaks at 16 and 27 min likely represent the two fragments generated by cleavage of proGRP31–125 after Pro78-Lys79. Similar expression and purification strategies were used for proGRP18–125 and proGRP42–98, and all three fragments were characterized by PAGE (Fig. 2BGo), N-terminal sequencing, and mass spectrometry. The observed masses for the proGRP fragments were all within 0.3% of the predicted values, and the protein yields obtained from 0.2 liters of bacterial culture ranged from 0.2 mg for proGRP18–125 to 1.0 mg for proGRP31–125.


Figure 2
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  		FIG. 2. Purification of proGRP31–125. Human proGRP31–125 was expressed in E. coli as a fusion protein with glutathione S-transferase. The fusion protein was purified from bacterial lysates by chromatography on glutathione-agarose and cleaved by treatment with thrombin, as described in Materials and Methods. The supernatant was separated from the agarose beads by centrifugation and subjected to reverse phase HPLC (A) as described in Materials and Methods. The major peak at 39 min detected by absorbance at 280 nm is glutathione S-transferase. The peak marked with an asterisk at 25 min contained proGRP31–125. HPLC-purified proGRP18–125, proGRP31–125, and proGRP42–98 were electrophoresed on 20% polyacrylamide gels with a buffer system designed for peptides and proteins in the molecular weight range 5–70 kDa (22 ) and visualized by staining with Coomassie blue (B). The bands observed migrated more slowly than expected, but the molecular weights determined by mass spectrometry were in excellent agreement with the predicted values. 1, Molecular weight markers; 2, proGRP18–125; 3, proGRP31–125; 4, proGRP42–98.

 
Recombinant C-terminal proGRP fragments are biologically active
As a first step to defining biologically active regions of the C-terminal proGRP fragments, the ability of proGRP18–125 to stimulate proliferation and migration of DLD-1 colorectal carcinoma cells was investigated. Treatment with proGRP18–125 at a concentration of 50 or 200 nM produced an increase in cell number as measured by the MTT colorimetric assay in DLD-1 cells (supplemental Fig. 1A, published as supplemental data on The Endocrine Society’s Journals Online Web site at http://endo.endojournals.org) and also increased cell migration measured in a wound healing assay (supplemental Fig. 1B). Although we have previously shown that DLD-1 cells express both GRP-R and the orphan receptor BRS-3 (7), the observation that the GRP-R antagonist [D-Phe6, Leu-NHet13, des-Met14]-bombesin (6, 7, 8, 9, 10, 11, 12, 13) at a concentration of 50 nM did not block stimulation of either proliferation or migration indicated that the GRP-R was not involved (supplemental Fig. 1).

To confirm that the biological activity of proGRP18–125 was independent of amino acids 18–27 and to narrow down the region within C-terminal proGRP responsible for the biological activity, the shorter fragments proGRP31–125 and proGRP42–98 were generated as described in Materials and Methods. Both peptides at a concentration of 10 nM significantly stimulated proliferation in the colorectal carcinoma cell line DLD-1 (Fig. 3AGo) and the prostate carcinoma cell line DU145 (Fig. 3BGo). The stimulation was not significantly reduced by inclusion of the GRP-R antagonist, which was able to block GRP-stimulated proliferation in DLD-1, DU145, and HT-29 cells (Fig. 3Go, A–C). Proliferation of the colorectal carcinoma cell line SW1222, endometrial carcinoma cell line Ishikawa, and embryonic kidney cell line HEK293 was not affected by GRP or proGRP31–125 or proGRP42–98 (data not shown). The fact that no binding of proGRP18–125 or proGRP31–125 was detected to cell lines stably transfected with either GRP-R or BRS-3 (data not shown) confirms that the biological activity of C-terminal proGRP fragments is exerted through receptors distinct from GRP-R and BRS-3.


Figure 3
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  		FIG. 3. Recombinant C-terminal proGRP fragments stimulate proliferation of DLD-1 and DU145 cells. Proliferation of a panel of cell lines in response to 10 nM GRP, proGRP31–125, or proGRP42–98, without and with the GRP-R receptor antagonist (D-Phe6,Leu-NHet13, des-Met14)-bombesin (6 7 8 9 10 11 12 13 14 ) (A), was assessed by a [3H]thymidine incorporation assay. The human cell lines tested were the colorectal carcinoma cell line DLD-1 (A), prostate carcinoma cell line DU145 (B), and colorectal carcinoma cell line HT29 (C). Data are the means ± SEM of at least three separate experiments each performed in quadruplicate and are expressed as a percentage of the control. Statistical significance was assessed by one-way ANOVA (*, P < 0.05; **, P < 0.01; #, P < 0.001, compared with control; {wedge}, P < 0.05, compared with GRP-stimulated sample). proGRP31–125 and proGRP42–98 significantly stimulated proliferation of DLD-1 and DU145 cells, and the stimulation was not inhibited by the GRP receptor antagonist.

 
Biological activity of fragments of proGRP42–98
Because proGRP42–98 was biologically active, further fragments were synthesized to define more precisely the active region of proGRP42–98. The synthetic peptides proGRP47–68 and [Tyr79]proGRP80–97 at a concentration of 100 nM significantly stimulated proliferation of DLD-1 colorectal carcinoma cells, and the extent of stimulation was not significantly different from that induced by recombinant proGRP42–98 at the same concentration (Fig. 4AGo). In titration experiments a maximum response was observed at 100 nM proGRP47–68 (184 ± 10%, Fig. 4BGo) or 10 nM [Tyr79]proGRP80–97 (215 ± 26%, Fig. 4CGo). No significant stimulation of proliferation was detected at concentrations of either peptide lower than 0.1 nM (Fig. 4Go, B and C). When the two peptides were tested together at a submaximal concentration of 0.1 nM, the response (151 ± 3%) was not significantly greater than that observed with either peptide alone at the same concentration (proGRP47–68, 141 ± 3%; [Tyr79]proGRP80–97, 162 ± 10%).


Figure 4
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  		FIG. 4. Synthetic C-terminal proGRP fragments stimulate proliferation and migration of DLD-1 cells. Proliferation of the human colorectal carcinoma cell line DLD-1 in response to 10 nM GRP or 100 nM recombinant proGRP42–98, synthetic proGRP47–68, or synthetic [Tyr79]proGRP80–97 (A), increasing concentrations of synthetic proGRP47–68 (B), or increasing concentrations of synthetic [Tyr79]proGRP80–97 (C) was assessed by a [3H]thymidine incorporation assay. In a typical proliferation assay, the radioactivity incorporated in control wells was 1122 ± 60 cpm, compared with 1840 ± 125 cpm in wells stimulated with 10 nM proGRP47–68 and 1774 ± 214 cpm in wells stimulated with 10 nM [Tyr79]proGRP80–97. D, Migration of DLD-1 cells in response to 10 nM proGRP47–68 or [Tyr79]proGRP80–97 was measured in a wound-healing assay. The width of the wound was measured at five distinct sites from the microphotographs taken at 17 h (gray bars) or 24 h (white bars) and expressed as a percentage of the value at 0 h (black bars). Data are the means ± SEM of at least three separate experiments each performed in triplicate. Statistical significance was assessed by one-way ANOVA (*, P < 0.05; **, P < 0.01; #, P < 0.001, compared with control). The synthetic fragments proGRP47–68 and [Tyr79]proGRP80–97 significantly stimulated proliferation and migration of DLD-1 cells.

 
The ability of the two proGRP fragments to stimulate migration was then measured in a wound-healing assay. Treatment of DLD-1 cells with 10 nM proGRP47–68 or proGRP80–97 caused a time-dependent decrease in wound size (Fig. 4DGo). After treatment with proGRP47–68 or 10 nM proGRP80–97 for 24 h, the wound size was 11 ± 3 and 7 ± 1%, respectively, whereas in the untreated control, the wound was still 32 ± 3% of its original size.

Activation of intracellular signaling pathways by proGRP fragments
Increases in inositol phosphate production and tyrosine phosphorylation occur soon after the binding of many peptides to G protein-coupled and growth factor receptors, respectively. To elucidate the intracellular signaling pathways activated by proGRP fragments inositol phosphate production was measured in DLD-1 colorectal carcinoma cells after treatment with proGRP47–68 or proGRP80–97. Either peptide at a concentration of 1 nM significantly stimulated inositol phosphate production (Fig. 5AGo). A higher concentration of 10 nM did not increase inositol phosphate production significantly, perhaps because of receptor down-regulation as previously reported in the case of the GRP-R (7). In contrast, neither peptide stimulated tyrosine phosphorylation in DLD-1 cells (data not shown).


Figure 5
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  		FIG. 5. Synthetic C-terminal proGRP fragments stimulate inositol phosphate production and MAPK kinase activity in DLD-1 cells. A, Production of inositol phosphates by DLD-1 human colorectal carcinoma cells after incubation with 1 or 10 nM synthetic proGRP47–68 or synthetic [Tyr79]proGRP80–97 was measured as described in Materials and Methods. In a typical experiment, the radioactivity incorporated in control wells was 4710 ± 387 cpm, compared with 6883 ± 314 cpm in wells stimulated with 1 nM proGRP47–68 and 6360 ± 531 cpm in wells stimulated with 1 nM [Tyr79]proGRP80–97. B, Phosphorylation of MAPK in the human colorectal carcinoma cell line DLD-1 in response to 10 nM recombinant proGRP42–98, synthetic proGRP47–68 or synthetic [Tyr79]proGRP80–97 was measured as described in Materials and Methods. Cell lysates were electrophoresed on SDS-PAGE and Western blotted. Membranes were then incubated with antibodies against either total or phosphorylated p42/p44 MAPK, and bound antibody was detected with alkaline phosphatase-coupled IgG. The Western blot is representative of three separate experiments. C, Band densities were determined by densitometric analysis and are presented as the ratio of densities of phosphorylated protein to total protein. Data are the means ± SEM of three separate experiments, and statistical significance was assessed by one-way ANOVA (*, P < 0.05; #, P < 0.001, compared with control). The synthetic fragments proGRP47–68 and [Tyr79]proGRP80–97 significantly stimulated inositol phosphate production by phospholipase C, and MAPK kinase activity, in DLD-1 cells.

 
The MAPKs are key components of the intracellular signaling pathways that control cell proliferation (25). To elucidate intracellular signaling pathways further, the activity of MAPK kinase was measured by Western blotting of DLD-1 colorectal carcinoma cell extracts after treatment with 10 nM proGRP42–98, proGRP47–68, or proGRP80–97. Phosphorylation of both the 42- and 44-kDa forms of MAPK was significantly stimulated, and the extent of stimulation was similar with each peptide (Fig. 5Go, B and C).

Characterization of binding sites for C-terminal proGRP peptides
Because C-terminal proGRP fragments acted independently of either GRP-R or BRS-3, we characterized the binding sites responsible for their bioactivity. Binding experiments were performed with the synthetic peptides proGRP47–68 and [Tyr79]proGRP80–97. Both peptides were radioactively labeled by the chloramine T method, and the monoiodinated derivatives were purified by reverse phase HPLC. Binding of the iodinated peptides to the human colorectal carcinoma cell line DLD-1 in the absence or presence of the appropriate unlabeled proGRP peptides at a concentration of 1 µM was then measured as described in Materials and Methods. DLD-1 cells bound 6.0 ± 0.4% of added 125I-labeled proGRP47–68 and 7.4 ± 0.1% of added 125I-labeled [Tyr79]proGRP80–97. Nonspecific binding expressed as a percentage of total bound radioactivity was 25.9 ± 1.6 and 7.4 ± 0.8% for 125I-labeled proGRP47–68 and 125I-labeled [Tyr79]proGRP80–97, respectively. No significant inhibition of specific binding was observed in the presence of the GRP-R antagonist. HPLC analysis of the cell supernatant indicated that more than 80% of 125I-labeled proGRP47–68 and 70% of 125I-labeled [Tyr79]proGRP80–97 (data not shown) remained intact at the end of the assay.

To determine the affinity of the C-terminal proGRP receptor for C-terminal proGRP fragments, binding experiments with DLD-1 cells were performed in the presence of increasing concentrations of unlabeled proGRP47–68 (Fig. 6AGo) and [Tyr79]proGRP80–97 (Fig. 6BGo). The data for proGRP47–68 were well fitted by a one-site model with dissociation constant (Kd) 170 ± 30 nM. In the case of proGRP80–97, a better fit was obtained with a two-site model with Kd1 40 ± 50 fM (32%) and Kd2 38 ± 12 nM (68%). The number of sites per cell was 4.8 ± 0.4 x 106 for proGRP47–68 and 1.5 ± 0.1 x 106 for [Tyr79]proGRP80–97. Similar binding properties were also observed with HCT-15 colorectal carcinoma cells (data not shown). The colorectal carcinoma cell line HT 29 and the endometrial cell line Ishikawa, which do not proliferate in response to proGRP42–98, express less than 20% of the number of binding sites found on DLD-1 cells (data not shown).

All of the other proGRP fragments tested significantly inhibited binding of iodinated proGRP47–68 (Fig. 6AGo). The most effective inhibitor observed was proGRP80–97 with Kd 90 ± 30 nM. The best-fit Kd values for proGRP31–125 and proGRP42–98 were 3.3 and 1.2 µM, respectively, but in both cases these values should be regarded as estimates only because the observed inhibition at 1 µM peptide was less than 50%. Similarly all of the other proGRP fragments tested significantly inhibited binding of iodinated [Tyr79]proGRP80–97 (Fig. 6BGo). The most effective inhibitor was proGRP47–68 with Kd 130 ± 40 nM. The Kd values for proGRP31–125 and proGRP42–98 were 2.2 and 4.7 µM, respectively, but in both cases these values should be regarded as estimates only because the observed inhibition at 1 µM peptide was less than 50%. Interestingly there was no indication of a second higher affinity binding site with any fragment other than proGRP80–97.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
High-molecular-mass peptides derived from proGRP are present, often at much higher concentrations than GRP18–27, in the tumors and circulation of patients with SCLC (26, 27, 28, 29, 30), medullary thyroid carcinoma (31, 32), metastatic or androgen-independent prostate carcinoma (33, 34), or colorectal carcinoma (9). Assays that measure proGRP31–98 have been developed for cancer diagnosis and treatment monitoring (19). There is also evidence for the presence of larger forms of GRP in ovine and human uterus and the circulation of both species (35, 36). Previously Miyake et al. (28) reported the expression of proGRP31–98 as a TrpE fusion protein from a chemically synthesized gene. The possibility that some of the above proGRP fragments might be biologically active has not been investigated so far, although the extensive homology between mammalian species in the C-terminal region of proGRP indicated that a functional role was likely. The above observations and our recent report that the GRP precursor proGRP1–125 was biologically active when expressed as a His-tagged fusion protein in E. coli (9), make a convincing argument that the regions within proGRP31–125 responsible for the bioactivity should be determined. In the present paper, we report the preparation of several recombinant proteins derived from the C-terminal portion of proGRP, the demonstration that they are biologically active, and the definition of the binding properties of the receptor responsible for the activity.

Recombinant proGRP18–125 and proGRP31–125 were first prepared and shown to be biologically active (supplemental Fig. 1). Although we have previously shown that DLD-1 cells express both GRP-R and the orphan receptor BRS-3 (7), the observation that a GRP-R antagonist was unable to block the effect of either peptide indicated that the GRP-R was not involved. This conclusion was confirmed by the observation that no binding of proGRP18–125 or proGRP31–125 was detected to cell lines stably transfected with either GRP-R or BRS-3 (data not shown).

To narrow down the region responsible for activity, proGRP42–98 was generated and its biological activity compared with proGRP31–125 in a panel of six cell lines. ProGRP31–125 and proGRP42–98 significantly stimulated proliferation of the colorectal carcinoma cell line DLD-1 (Fig. 3AGo) and the prostate carcinoma cell line DU145 (Fig. 3BGo) but not the colorectal carcinoma cell line HT29 (Fig. 3CGo). As expected, the stimulation was unaffected by the inclusion of a GRP-R antagonist in the assay. In contrast, GRP18–27 significantly stimulated proliferation of DLD-1, DU145, and HT29 cells (Fig. 3Go, A–C), and in all three cases the stimulation was significantly reduced in the presence of the GRP-R antagonist. Interestingly all three cell lines produce proGRP (33–94 fmol per 106 cells) but no detectable amidated GRP (9), so the possibility of an autocrine growth loop should be borne in mind. Proliferation of the colorectal carcinoma cell line SW1222, endometrial carcinoma cell line Ishikawa, and embryonic kidney cell line HEK293 was not affected by GRP or proGRP31–125 or proGRP42–98 (data not shown). Taken together, these results indicate that the biological effects of C-terminal proGRP fragments are not unique to DLD-1 cells and confirm the conclusion that C-terminal proGRP fragments act independently of the GRP receptor.

To further delineate the region within proGRP42–98 responsible for bioactivity, the peptides proGRP47–68 and [Tyr79]proGRP80–97 were synthesized. These sequences were based on the two assumptions that the most highly conserved regions within the proGRP42–98 sequence would be responsible for receptor binding and that any processing of proGRP42–98 would likely occur after lysine residues. Searching of vertebrate nucleotide and protein sequence databases did not reveal any close relatives of the two peptides, apart from proGRP from species other than humans. Both peptides were independently able to stimulate proliferation (Fig. 4AGo), migration (Fig. 4DGo), inositol phosphate production (Fig. 5AGo), and MAPK kinase activation (Fig. 5Go, B and C) in DLD-1 cells.

Because C-terminal proGRP fragments do not act through the known receptors BRS-3 and GRP-R, the receptor through which these peptides act was characterized. Radiolabeled proGRP47–68 and [Tyr79]proGRP80–97 both bound to DLD-1 (Fig. 6Go) and HCT-15 cells (data not shown), and binding of neither peptide was inhibited by a GRP-R antagonist. The binding data from titration experiments with proGRP47–68 were well fitted by a one-site model with Kd 170 ± 30 nM (Fig. 6AGo). In the case of proGRP80–97, a better fit was obtained with a two-site model with Kd1 40 ± 50 fM (32%) and Kd2 38 ± 12 nM (68%) (Fig. 6BGo). In contrast, both proGRP47–68 and [Tyr79]proGRP80–97 stimulated DLD-1 cell proliferation at concentrations between 0.1 and 100 nM (Fig. 4Go, B and C). One possible explanation for the apparent discrepancy between the proliferation and binding data are that fractional occupancy of the receptor is sufficient to stimulate proliferation maximally. For example, in the case of the cholecystokinin receptor, 90% maximal amylase release is achieved at approximately 10% receptor occupancy (37).

The binding site possesses some other unusual features. First, the observation that each short peptide was able to compete with the other for binding suggests that the binding sites for proGRP47–68 and proGRP80–97 overlap to some extent. The fact that the biological activities of the two peptides are not additive is consistent with competitive binding. Second, the N- and C-terminal extensions of the two longer peptides may interfere with access to the receptor binding pocket. The data on biological potency suggest a reduced affinity for longer peptides because stimulation of DLD-1 cell proliferation by proGRP47–68 and [Tyr79]proGRP80–97 was nearly maximal at 10 nM (Fig. 4Go, B and C), but stimulation by proGRP18–125 was detected only in the range 50–200 nM (supplemental Fig. 1A). The observation that binding of either proGRP47–68 or proGRP80–97 was only partially inhibited by proGRP31–125 or proGRP42–98 at 1 µM (Fig. 6Go) is consistent with this conclusion. Our current model for the receptor binding site therefore envisions a restricted pocket capable of accommodating either proGRP47–68 or proGRP80–97 but with some overlap hindering the binding of both short peptides simultaneously. Our data also raise the possibility that the two divergent receptor binding domains of proGRP47–68 and proGRP80–97 adopt identical conformations similar to the case of PTH and PTHrP. For both PTH and PTHrP, the region 15–34 in which only three amino acids are identical functions as the principal PTH/PTHrP receptor binding domain. The observation that peptides based on the sequences PTH14–34 and PTHrP14–38 compete equally for binding with radiolabeled PTH1–34 or PTHrP1–36 to the PTH/PTHrP receptor (38, 39) indicates that two peptides with virtually no sequence homology can bind to a common site on the same receptor. Other models incorporating allosteric interactions between distant sites are also possible. In any case the data are consistent with the hypothesis that DLD-1 cells express a receptor, distinct from GRP-R, which recognizes two independent epitopes within the proGRP42–98 sequence.

The question then arises whether peptides such as proGRP47–68 and proGRP80–97 occur naturally. The presence of several posttranslational processing products of proGRP has been reported previously (18). In particular, in culture media from cell lines derived from SCLC tumors cleavage occurred between Lys79 and Ala80 (40). In the SCLC cell line H345, cleavages also occurred between Arg71 and Asn72, Glu40 and Arg41, and Gly63 and Leu64 (41). Combinations of these cleavages would generate a variety of proGRP fragments including proGRP41–63 and proGRP80–125. More precise definition of stored and circulating fragments derived from the C terminus of proGRP will require the development of region-specific antibodies capable of recognizing epitopes within the proGRP31–125 sequence.

The results in the present paper may be summarized as follows. Peptides derived from the C terminus of proGRP stimulate both proliferation and migration of the human colorectal carcinoma cell line DLD-1. The failure of these peptides to bind to either the orphan receptor BRS-3 or the GRP-R clearly demonstrates that the effects of the proGRP-derived peptides are mediated through a novel receptor. A candidate binding site that recognizes both N- and C-terminal epitopes within the proGRP42–98 sequence has been defined. Future work should be directed toward characterization of the structure of the receptor for these peptides and toward determining to what extent these peptides are generated in tumors derived from lung, prostate, or colorectal tissue. In the longer term, such studies would permit the design of antagonists that might prove useful as cancer therapies.


   Acknowledgments
 
We thank Professor James F. Battey (National Institute for Deafness and Other Communication Disorders, National Institutes of Health, Bethesda, MD) for generously providing the DNA encoding human PreproGRP1–125 cloned into the pSp64 vector and the BALB 3T3 cell lines expressing the GRP receptors, GRP-R and BRS-3. We thank Dr. Mustafa Ayan from the Baker Heart Research Institute (Prahran, Victoria, Australia) for the mass spectroscopy.


   Footnotes
 
This work was supported in part by grants from the National Health and Medical Research Council of Australia and the Austin Hospital Medical Research Foundation.

Disclosure Summary: O.P., C.D., A.S., and G.S.B. have nothing to declare.

First Published Online November 22, 2006

Abbreviations: BRS-3, Bombesin receptor subtype 3; FBS, fetal bovine serum; GRP, gastrin-releasing peptide; GRPgly, glycine-extended GRP; GRP-R, GRP receptor; Kd, dissociation constant; MTT, 3-[4,5-dimethylthiazol-2-yl]2,5-diphenyltetrazolium bromide; NMB, neuromedin B; SCLC, small cell lung carcinoma.

Received April 13, 2006.

Accepted for publication November 10, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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