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Endocrinology Vol. 139, No. 10 4329-4336
Copyright © 1998 by The Endocrine Society

ARTICLES

Glucose Stimulation of Pancreatic ß-Cell Lines Induces Expression and Secretion of Dynorphin

Knud Josefsen, Karsten Buschard, Lone Reinholdt Sørensen, Michael Wøllike, Rolf Ekman and Mark Birkenbach

Bartholin Instituttet (K.J., K.B., L.R.S., M.W.), Kommunehospitalet, DK-1399 Copenhagen K, Denmark; Department of Clinical Neuroscience (R.E.), Göteborg Universitet, Mölndal Hospital, S431-80 Sweden; and Kovler Viral Oncology Laboratory (M.B.), University of Chicago, Chicago, Illinois 60637

Address all correspondence and requests for reprints to: Knud Josefsen, M.D., Ph.D., Bartholin Instituttet, Kommunehospitalet, DK-1399 Copenhagen K, Denmark. E-mail: josefsen{at}dadlnet.dk


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
To investigate adaptive responses of pancreatic ß-cells to hyperglycemia, genes induced by glucose stimulation were identified by subtraction cloning. Among 53 clones representing differentially expressed genes, 20 encoded the endogenous opioid precursor, prodynorphin. The amino acid sequence of murine prodynorphin is identical to the rat protein in sequences comprising the opioid peptides and 86% identical in the remainder of the molecule. Stimulation of MIN6 cells increased prodynorphin RNA levels to more than 20-fold in proportion to physiological glucose concentrations. Similar induction levels were observed in murine ßTC3 and rat Rinm5F ß-cell lines. Prodynorphin RNA expression increased within 1 h of glucose stimulation, achieved maximal levels by 4 h, and remained elevated for at least 24 h. By using RIA, MIN6 cells were shown to contain and secrete increased amounts of dynorphin-A following glucose stimulation. Treatment of MIN6 cells with KCl, forskolin, or isobutyl-methyl-xanthine strongly induced prodynorphin RNA expression, suggesting that induction may be related to secretion-coupled signaling pathways. The induction of prodynorphin in several ß-cell lines is consistent with previous demonstrations of ß-cell synthesis of other endogenous opioids, including ß-endorphin, and suggests that opioids may have a potentially significant role in regulating ß-cell secretion.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE ß-CELLS of pancreatic islets of Langerhans modulate serum glucose levels through precisely regulated release of insulin from preformed granules. A sequence of rapid electrophysiologic events initiated by glucose phosphorylation culminates in depolarization of ß-cell membranes and opening of voltage-sensitive, L-type Ca++-channels (1). The resulting increase in intracellular Ca++-ion concentration activates exocytosis of insulin granules.

Chronic glucose stimulation may alter ß-cell responses to glucose in vivo and in vitro (reviewed in Ref. 2). In vivo, short-term exposure to glucose can either augment or inhibit insulin secretion in response to subsequent stimulation (3). In contrast, prolonged hyperglycemia typically reduces ß-cell responsiveness. It has been suggested that the latter phenomenon, referred to as glucose toxicity, may relate to the impaired ß-cell glucose responses that characterize noninsulin-dependent diabetes mellitus (4, 5, 6).

Short-term glucose effects on ß-cell responsiveness may be mediated in part through actions of autocrine factors. In resting ß-cells, insulin is stored in preformed granules that also contain chromogranin A, the prohormone convertases 2 and 3 (PC2 and PC3), and amylin (also known as islet amyloid polypeptide, IAPP) (7). A direct inhibitory effect of amylin on insulin secretion by perfused rat islets in vitro has been demonstrated (reviewed in Ref. 8). Pancreastatin, derived by proteolytic cleavage of chromogranin A, also inhibits insulin secretion in vitro (9). Lastly, neuropeptide Y, which is synthesized and secreted by both primary rat islet cells and many rodent ß-cell lines, potently inhibits glucose-stimulated insulin secretion (10). Together these data suggest that a variety of peptide factors produced by ß-cells and cosecreted with insulin may exert important autocrine regulatory effects that could alter responses to sequential or prolonged stimuli.

Long-term effects of glucose are likely mediated largely through changes in gene expression. In vitro, chronic stimulation of HIT-T15 cells results in diminished insulin gene transcription and progressive reduction in insulin release. This has been attributed to loss of essential transcriptional activators, somatostatin transcription factor-1 (IPF-1) and RIPE3b1 (6). In vivo, several critical ß-cell genes, including insulin, undergo transcriptional changes during glucose stimulation (11). Moreover, expression of the putative autocrine factor amylin has been shown to be glucose regulated (12). Lastly, chronic hyperglycemia has been associated with reduced transcription of IPF-1 and the GLUT-2 glucose transproter (13). These findings suggest that glucose-induced changes in ß-cell gene transcription may alter responses to subsequent stimulation.

The present study was undertaken to examine the effects of glucose on ß-cell gene expression. The foregoing data indicate that glucose-regulated genes may have important roles as modulators of ß-cell responses to glucose. In addition, recent evidence suggests that expression of some antigenic targets of autoimmune responses in type 1 insulin-dependent diabetes may also be glucose-regulated (14). Using subtractive hybridization, we have identified several genes induced by glucose in the murine ß-cell line, MIN6. Among these are genes encoding glycolytic pathway enzymes, three immediate-early response transcription factors (14A ) and the endogenous opioid precursor, prodynorphin. We show that prodynorphin RNA expression is induced by glucose stimulation of several ß-cell lines and that synthesis and secretion of dynorphin peptides are intimately associated with insulin release in MIN6 cells. Because endogenous peptides and their receptors have been identified in pancreas (15) and modulate islet transcription (16), prodynorphin-derived peptides from islets could participate in carbohydrate regulation through paracrine or endocrine actions.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Tissue culture
The SV40-transformed murine ß-cell line, ßTC3 (17), and early passage (<40) MIN6 cells (18) were cultured in 90% DMEM (22.5 mM glucose), 10% FBS (Life Technologies, Gaithersburg, MD), with three medium changes per week. Hamster HIT-T15 (19) and rat RINm5F cells (20) were grown in 90% RPMI-1640 (11 mM glucose), 10% FBS. For stimulation, cells were first cultivated for 24 h in 90% DMEM, 10% FBS containing 1 mM (MIN6), 0.5 mM (ßTC3), 0.1 mM (HIT-T15) or 0.01 mM (RINm5F) glucose. Glucose (20 mM) was subsequently added to the cultures for 24 h or the cells were left unchanged (unstimulated controls) for a similar time.

Islets of Langerhans
Male Lewis rats, aged 60 days, were purchased from Møllegrd, Ll. Skensved, Denmark and allowed free access to food and water. Islets were isolated as previously described (21). All animal work was carried out under the regulations set by the Animal Experiments Inspectorate.

RIA
Aliquots of culture supernatants were cleared by centrifugation at 200 x g and frozen at -70 C for later determination of insulin and dynorphin concentrations by RIA. For insulin measurements (22), complexes of insulin and guinea pig antiinsulin antibody (1:36,000; Novo Nordisk, Bagsværd, Denmark) were precipitated with rabbit antiguinea pig secondary antibody (Dako, Glostrup, Denmark), using purified rat insulin (Novo Nordisk) as a standard. Immunoreactive dynorphin A concentrations were determined using a monospecific rabbit antiserum at a final dilution of 1:12,500 (23). This antiserum reacts strongly with dynorphin A(1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17) and (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13) but has a 30-fold lower affinity for dynorphin A(1, 2, 3, 4, 5, 6, 7, 8, 9), (1, 2, 3, 4, 5, 6, 7, 8, 9, 10), and (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11) (23). Extraction from cells was performed by boiling for 10 min in 0.5 M acetic acid, followed by ultrasonication for 30 seconds and centrifugation. The pellet was reboiled in 0.9% NaCl, ultrasonicated, and spun. The supernatants were combined and lyophilized.

Subtraction cloning
Complementary DNA (cDNA) clones of genes expressed in glucose-stimulated cells at higher levels than in unstimulated cells were identified as previously described (24). In brief, total cellular RNA was isolated from stimulated and control MIN6 cells by the acid guanidinium thiocyanate-phenol-chloroform extraction method (25), and polyadenylated fractions were selected chromatographically using oligo-deoxythymidylate-cellulose. 32P-labeled cDNA probes were synthesized by reverse transcription of RNA from stimulated cells and combined with 20 µg photobiotinylated RNA from unstimulated cells. Hybrids were removed by addition of streptavidin followed by extraction with neutral phenol-chloroform. Remaining radiolabeled, single-stranded cDNA was used as a probe directly to screen a lambda gt10 bacteriophage cDNA library prepared from glucose-stimulated MIN6 cells as previously described (24).

RNA analysis
Prodynorphin and ß-actin messenger RNA (mRNA) expression were evaluated by blot hybridization. Total cellular RNA (7–10 µg per sample) was fractionated on 1% agarose-formaldehyde gel and transferred to activated nylon membranes (GeneScreen Plus, New England Nuclear, Billerica, MA). Radioactive 32P-labeled probes were synthesized by random hexamer nucleotide priming, purified by gel exclusion chromatography and hybridized to RNA blots in hybridization buffer (50% formamide; 6 x SSPE; 1 x Denhardt; 1% SDS; 100 µg/ml herring testis DNA). Filters were washed three times at 62 C in 0.2 x SSC, 1% SDS, and exposed to x-ray film (Reflection, New England Nuclear, Boston, MA) at -70 C. Autoradiographic signals were quantified densitometrically using a flatbed scanner (Hewlett-Packard Scanjet 4c/T; Greeley, CO) and SigmaGel software (Jandel Scientific, San Rafael, CA). This method was linear over at least one decade.

Sequencing
DNA nucleotide sequences were determined by the dideoxynucleotide chain termination method using T7 DNA polymerase (Pharmacia, Uppsala, Sweden) and specific synthetic oligonucleotide primers. Compressions and sequence ambiguities were resolved by bidirectional sequencing using 7-deaza dGTP nucleotides. Homology comparisons with database sequences were done using the BLAST Network Service at the National Center for Biotechnology Information (NCBI).

Statistics
Student’s t test was used for evaluating differences between means. Values of P < 0.05 were considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Isolation of glucose-induced genes
A cDNA library prepared from glucose-stimulated murine MIN6 ß-cell RNA was screened using 32P-labeled subtracted cDNA probes specific for genes expressed at higher levels in stimulated than in unstimulated cells. From a total of 66,000 recombinant phage clones screened, 53 clones of differentially expressed genes were isolated and subcloned for further analysis. Nucleotide sequences of cDNA insert ends and restriction digestion analysis indicated that 20 of these were derived from the same gene. Comparison of partial sequences from these clones with nucleotide sequence databases of the National Center for Biotechnology Information indicated a high degree of similarity with a previously identified rat gene encoding the endogenous opioid precursor, prodynorphin (Preproenkephalin B; GenBank M32784, M10088). Other genes isolated encoded glycolytic enzymes, including: triose phosphate isomerase; phosphoglycerate kinase; lactate dehydrogenase; and enolase (Josefsen, K., N. Grunnet, K. Buschard, and M. Birkenbach, manuscript in preparation). A single clone encoding the zinc-finger transcription factor egr-1 (Krox-24) was also isolated (14A ).

The largest prodynorphin clone contained a 2400 bp cDNA insert. Its complete nucleotide sequence and deduced amino acid translation product are compared with rat prodynorphin sequences in Fig. 1Go. The murine gene contains a single long open reading frame that begins with a consensus translational initiation methionine codon at nucleotide 204 and ends with a stop codon at nucleotide 948. These correspond precisely with positions of predicted translational initiation and termination codons of the rat prodynorphin gene. Within its open reading frame the murine gene shares 92% nucleotide identity with the rat gene. The deduced sequence of the primary murine translation product is 88% identical to the rat protein. Significantly, bioactive endoproteolytic cleavage products of the murine precursor polypeptide, which include dynorphins-A and -B, {alpha}- and ß-neo-endorphins, and leumorphin are predicted to be 100% identical to the corresponding rat peptides. A number of insertions and deletions in 3' and 5' untranslated regions distinguish the mouse gene from the rat gene such that the overall nucleotide identity is 86%.



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  		Figure 1. Complete nucleotide sequence of the mouse prodynorphin cDNA (GenBank accession no. AF026537). The predicted amino acid sequence is indicated by three-letter designation above each codon. The peptides in bold are {alpha}-neoendorphin (699–725), dynorphin-A (807–857), and dynorphin-B (864–947). Nucleotide and amino acid differences from the rat sequence are shown below the murine sequence. No amino acid differences between murine and rat sequences are identified in the predicted major opioid peptides. The poly-A adenylation site is indicated by underline.

 
Prodynorphin mRNA induction
The effects of glucose on prodynorphin expression were evaluated by blot hybridization of RNA extracted from stimulated and unstimulated cultures of various ß-cell lines. Following 24 h glucose deprivation, MIN6 cells were cultivated for 24 h in fresh medium containing either 20 mM (stimulated) or 1 mM glucose (unstimulated). Insulin concentrations were typically 6-fold higher in supernatants of glucose-stimulated MIN6 cells than in resting cells (9738 ± 730 ng/ml vs. 1613 ± 215 ng/ml, N = 4, P < 0.001). Comparable results were observed following stimulation of ßTC3 cells, demonstrating that this procedure efficiently stimulates the cells. Hybridization of 32P-labeled prodynorphin cDNA insert to RNA blots detected a 2.6-kb transcript that was much more abundant in glucose-stimulated MIN6, Rinm5F and ßTC3 cells than in the respective unstimulated cell preparations (Fig. 2Go). Densitometric quantification of autoradiograph band intensities indicated a 9-fold increase in steady-state prodynorphin RNA levels in MIN6 cells following glucose-stimulation, and more than 20-fold increases in Rinm5F and ßTC3 cells. Prodynorphin hybridization signals were much weaker in blots containing total cellular RNA (10 µg) extracted from hamster HIT-T15 ß-cells and failed to show increased expression following glucose stimulation. However, these cells have consistently exhibited poor glucose response characteristics in our laboratory as well as in others (26). Surprisingly, prodynorphin RNA was also undetectable by blot hybridization in unfractionated cells of primary rat islets of Langerhans cultivated in vitro (data not shown). This failure is likely due to sensitivity limitations of this technique because we (see below) and others (15, 27) have consistently detected expression of dynorphin peptides in rat islet cells. Alternatively, prodynorphin RNA may be labile and rapidly degraded in islet cells cultivated in vitro, while prodynorphin peptides persist for an extended time in a stably stored form.



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  		Figure 2. Prodynorphin mRNA induction in MIN6, ßTC3 and Rinm5F cells. Cells were grown in high (H) or low (L) glucose medium for 24 h following 24 h culture in low glucose medium. Stimulatory glucose concentrations were 20 mM glucose. Nonstimulatory concentrations were 1 mM (MIN6), 0.5 mM (ßTC3), or 0.01 mM (RINm5F) glucose. Lower panel shows ß-actin control hybridizations.

 
The magnitude of maximal prodynorphin mRNA induction was graduated in proportion to stimulatory glucose concentrations. Hybridization of RNA from MIN6 cells stimulated with 0 to 20 mM glucose demonstrated a threshold for prodynorphin RNA induction at a concentration of 3 mM (Fig. 3Go). Maximal stimulation was achieved at 10 mM glucose that induced prodynorphin RNA more than 20-fold over unstimulated cells. Glucose concentrations above this level failed to increase prodynorphin RNA expression further or actually diminished the magnitude of induction. Densitometric scanning indicated that prodynorphin hybridization signal intensities closely matched a power function of the glucose concentration (exponent approximately 1.5) over the physiological range from 4–10 mM.



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  		Figure 3. Dependence of prodynorphin mRNA induction in MIN6 cells on stimulatory glucose concentration. Cells were cultivated in low glucose medium for 24 h and subsequently exposed for 24 h to media containing the indicated glucose concentrations. Based on the ß-actin hybridization (center) the induction ratio was calculated (bottom). The ratio expresses induction relative to the level of prodynorphin in unstimulated cells.

 
To determine the time course of prodynorphin induction, RNA was isolated from MIN6 cells after glucose stimulation of varying duration (Fig. 4Go). An increase in steady state prodynorphin RNA first became evident by 60 min post stimulation, with maximal levels reached at 4 h. This contrasts with the kinetics of egr-1 RNA accumulation described previously (14A ) which peaks by 30 min after exposure to glucose.



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  		Figure 4. Time course of prodynorphin mRNA induction in glucose-stimulated MIN6 cells. Forty-eight hours glucose-deprived cells were incubated in high glucose medium (20 mM glucose) and RNA isolated at the indicated times after stimulation. ß-actin control hybridization is shown at the bottom.

 
Secretion of dynorphin
Concentration of the endoproteolytic cleavage product of prodynorphin, dynorphin-A, was measured by RIA in culture supernatants and cytoplasmic lysates. Following 24 h incubation in high glucose medium, dynorphin-A content in MIN6 cells increased from 249 ± 42 to 487 ± 38 pmol/107 cell (P < 0.001, N = 12). These results confirm that induction of prodynorphin RNA is accompanied by a corresponding increase in synthesis or processing of prodynorphin protein. Simultaneously, concentrations of dynorphin-A in MIN6 culture supernatants increased from 25 ± 7.1 to 34.5 ± 9.1 pM (P = 0.04, N = 8). These values are significantly higher than the basal dynorphin-A concentration of 16.4 pM (N = 5) measured in cell-free culture medium. Together these results indicate that MIN6 cells constitutively release small amounts of the peptide, and secretion of dynorphin A is significantly augmented by glucose. The effects of glucose on dynorphin-A synthesis and release were potentiated by stimulation of protein kinase A or C with cAMP agonists or phorbol ester, respectively (data not shown).

Synthesis and release of dynorphin-A by in vitro cultivated rat islets of Langerhans were also assayed. Expression of dynorphin was detected in both stimulated (106 ± 80 pmol/107cells, N = 5) and unstimulated (108 ± 80 pmol/107 cells) islets at similar concentrations. These levels were considerably higher than baseline concentrations measured in the medium used for stimulation (16.4 pM), and corroborate previous immunohistochemical and receptor ligand displacement studies from other laboratories demonstrating dynorphin peptide expression in rat islets (15, 27). Our results also indicated that islet dynorphin-A concentrations were not significantly affected by glucose (P = 0.97). Because the islet cell populations used were not fractionated and since previous studies have detected dynorphin peptides predominantly in {alpha}-cells (15, 27), it is likely that most of the dynorphin-A detected in our assays is {alpha}-cell derived and therefore unresponsive to glucose stimulation. Augmented dynorphin-A synthesis in ß-cells may be further masked by a corresponding increase in secretion rate. The concentration of dynorphin in supernatants of both unstimulated (21.1 ± 7.8 pM) and 24 h glucose-stimulated (26.6 ± 10.2 pM, N = 11) islet cultures was also assayed and was significantly higher than levels measured in fresh culture media (16.4 pM), indicating that dynorphin is likely released constitutively from islets. However, the small increase in dynorphin concentration observed following glucose stimulation failed to meet statistical significance (P = 0.13). Because dynorphin-A is rapidly degraded in tissue culture media with a half-life of less than 10 min (unpublished observations), it is likely that these numbers substantially underestimate quantities actually released. In sum these results demonstrate that dynorphin-A is present in and secreted by rat islet cells in vitro, but fail to prove that ß-cells specifically contribute to this process.

Specificity of dynorphin response
To exclude that prodynorphin induction could be part of a general growth response of ß-cells to glucose, the effects of serum stimulation on MIN6 prodynorphin expression were investigated. Following 24 h glucose- or serum-starvation, MIN6 cells were stimulated for 24 h with glucose or serum, respectively (Fig. 5Go). Glucose induced prodynorphin mRNA 5-fold when incubated in medium with standard serum content (10%), and 2.5-fold in low serum (0.5%) medium. It should be noted that under the latter conditions, the calculated induction ratio appears to be reduced due to elevated prodynorphin mRNA content in resting cells. In contrast, serum stimulation of serum-starved MIN6 cells did only slightly affect prodynorphin mRNA levels in the presence of either constant high (11 mM) or low (1 mM) glucose concentrations.



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  		Figure 5. Effects of serum and glucose on MIN6 expression of prodynorphin mRNA. MIN6 cells were glucose-stimulated for 24 h in low (0.5%, lanes 1 and 2) and normal (10%, lanes 5 and 6) serum following 24 h glucose deprivation. Under both stimulatory conditions prodynorphin mRNA is highly induced. Alternatively, cells were serum-stimulated in low- (1 mM, lanes 3 and 4), or high-glucose medium (11 mM, lanes 7 and 8) resulting in minimal changes in prodynorphin RNA levels.

 
The effects of various insulin secretagogues on prodynorphin RNA expression were investigated (Fig. 6Go). Forskolin and IBMX, stimulators of the cAMP-system, individually induced prodynorphin mRNA in MIN6 cells to levels higher than glucose alone. Moreover, both agents acted synergistically in combination with glucose to activate prodynorphin expression. Treatment of MIN6 cells with potassium chloride (KCl) also induced prodynorphin mRNA, indicating that membrane depolarization alone is sufficient to activate expression. The Ca++-ionophore, ionomycin, failed to alter prodynorphin RNA levels, suggesting that induction may require activation signals in addition to Ca++ influx. Insulin did not alter prodynorphin expression, indicating that direct effects of glucose on ß-cell activation pathways, rather than autocrine actions of secreted insulin, are responsible for prodynorphin mRNA induction. Finally, interleukins IL-1{alpha} and IL-1ß did not induce the gene above basal levels.



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  		Figure 6. Effects of insulin secretagogues on prodynorphin RNA expression in MIN6 cells. Following incubation for 24 h in low glucose medium cells were exposed to the indicated agents for 24 h at the following concentrations: 20 mM glucose; 0.3 U/ml insulin; 100 µM IBMX; 10 µM forskolin; 25 mM KCl; 1 µM ionomycin; 10 ng/ml interleukin-1{alpha}; 10 ng/ml interleukin-1ß.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The present study demonstrates that glucose stimulation induces expression of prodynorphin mRNA in the independently derived murine ß-cell lines MIN6 and ßTC3, and rat RinM5F ß-cells. An increase in both synthesis and secretion of the endoproteolytic cleavage product, dynorphin-A, was also observed following glucose stimulation of MIN6 cells. Expression of dynorphin-A was detected in primary islets, although glucose induction in ß-cells specifically could not be demonstrated. Dynorphin-A secretion by glucose-treated MIN6 cells is consistent with a model in which dynorphin peptides are stored with insulin in secretory vesicles and cosecreted following ß-cell stimulation. Our results are in agreement with previous reports describing expression of prodynorphin RNA and secretion of dynorphin peptides by ßTC3 cells (28), though regulation of prodynorphin expression by glucose was not examined in those studies. Significantly, these authors did not detect prodynorphin expression in any other cell line, including several neuroendocrine lines. Though the ß-cell lines used in our studies are transformed and capable of unlimited growth in vitro, they share many characteristics with primary ß-cells. MIN6 cells, in particular, closely resemble primary cells in their responses to glucose at physiologic concentrations, expression of prohormone convertases, PC2 and PC3, and processing of insulin (29). Together these findings argue strongly that the observed induction of prodynorphin expression by glucose is not an idiosyncratic phenomenon unique to a particular cell line but rather may be characteristic of ß-cell lines in general.

Prodynorphin, also known as proenkephalin-B, is one of three precursors of endogenous opioid peptides, the others being POMC and proenkephalin-A (reviewed in Ref. 30). Enzymatic cleavage of prodynorphin at dibasic motifs generates three bioactive peptides: dynorphin-A-(1–17), leumorphin, and ß-neo-endorphin (reviewed in Ref. 30). Each of these comprises the[Leu5]-enkephalin pentapeptide (L-enkephalin) sequence at its amino terminus. Additional peptides are produced in a highly tissue specific manner by subsequent proteolytic processing at various single basic residues or by cleavage at alternative dibasic sites. These peptides include dynorphin-32, dynorphin-A-(1–8), dynorphin-B-(1–13)(rimorphin) and {alpha}-neo-endorphin.

Prodynorphin is primarily synthesized in the brain in a variety of locations, including the hypothalamus, hippocampus, and striatum, and in spinal cord neurons of sensory receptive fields. In spinal posterior horn neurons, expression of prodynorphin-derived peptides is induced by noxious stimuli. A possible role in modulating nociception has been suggested (31, 32). Prodynorphin is also synthesized in endocrine organs such as the pituitary and adrenal glands, and in gonadal tissues. In the rat neurohypophysis, dynorphin peptides are copackaged with vasopressin into neurosecretory vesicles (33) and play a role in regulation of oxytocin secretion (34). Prodynorphin-derived peptides have also been detected in neural tissue associated with rat duodenum (35) and guinea pig ileum (36), from which it is secreted during peristalsis (37). Its function in these latter locations is not known. Little is known regarding the enzymes that process prodynorphin. However, recent data indicate that the serine protease family prohormone convertases, PC2 and PC3 (also known as PC1) may play an important role (38). PC2 and PC3 specifically cleave polypeptides at dibasic motifs. Expression of PC2 and PC3 in vivo is restricted to neuroendocrine cells, including pancreatic ß-cells and cells of the pituitary gland (reviewed in Ref. 39). Both proteases are also expressed in MIN6 cells (29). In ß-cells, these enzymes are essential for endoproteolytic processing of proinsulin and are packaged with insulin into secretory granules (reviewed in Ref. 39). PC3 has been shown to cleave prodynorphin into high molecular weight intermediate polypeptides (38). Similarly, PC2 and PC3 acting in concert can process POMC correctly in vitro (40). Together, these results suggest that the appropriate biochemical machinery to process prodynorphin into bioactive opioid peptides may be contained within ß-cell insulin secretory granules.

Biological effects of dynorphins are mediated by binding primarily to opioid receptors of the {kappa}-type, of which several subtypes exist (41). Lower affinity binding to µ- and {delta}-receptors may also take place (42). Binding of dynorphins to opioid receptors on cultured rat neurons causes inhibition of adenylate cyclase (43, 44) and closure of Ca++-channels (45).

Considerable evidence indicates that endogenous opioid peptides are expressed in endocrine pancreas. ß-endorphin, a POMC-derived peptide, has been consistently detected by immunohistochemical staining in glucagon-secreting pancreatic {alpha}-cells of rat, guinea pig and rabbit, as well as in rabbit ß-cells (15). Similarly, [Met5]-enkephalin, a proenkephalin-A-derived peptide, has been detected in islets of Langerhans in rat (46), human (47) and guinea pig (48). In contrast, studies on pancreatic expression of prodynorphin have yielded conflicting results and indicate marked variations among species. L-enkephalin was found immunohistochemically in both {alpha}- and ß-cells of rat islets (46), and by specific ligand displacement studies in human islets (47). Interpretation of these results, however, is complicated by the fact that L-enkephalin-related sequences are also present at carboxyl-termini of some proenkephalin-A peptides (30). Dynorphin-A immunoreactive peptides, which are exclusively derived from prodynorphin, have been identified in {alpha}-cells of rat pancreas (15, 27) but not in human (27) or guinea pig (48, 49) islets. However, prodynorphin peptides have been demonstrated in nonislet enterochromaffin cells of guinea pig pancreas (48, 49) and are the only endogenous opioids detectable in these cells.

In both rat {alpha}-cells and guinea pig enterochromaffin cells, prodynorphin peptides have been localized ultrastructurally to neurosecretory granules (15, 49). This localization pattern is analogous to that observed in the neurohypophysis and strongly suggests that dynorphin peptides could be cosecreted with glucagon upon stimulation of rat {alpha}-cells.

Transcription of the prodynorphin gene is largely regulated by promoter AP-1 and CREB binding sites (50). The multiple CREB elements are likely responsible for the stimulatory effects of cAMP agonists on prodynorphin transcription (51) as in ßTC3 cells following stimulation with 8-Br-cAMP (28). Interestingly, activation of CREB following glucose stimulation of mesangial cells has been demonstrated (52) and suggests a possible mechanism for prodynorphin induction by glucose in MIN6 cells. Regulation of prodynorphin transcription by AP-1 factors has been suggested by the temporal relation between AP-1 factor synthesis and subsequent induction of prodynorphin transcription in brain and spinal cord neurons following stimulation (53). We have previously shown that both c-fos and junB are up-regulated following glucose stimulation of MIN6 cells (14A ), suggesting that AP-1 factors may also contribute to prodynorphin induction in ß-cells.

Our observation that prodynorphin mRNA is induced by glucose in most ß-cell lines contrasts with results from immunolocalization studies that have consistently failed to detect dynorphin immunoreactivity in rodent or human ß-cells in vivo (15, 27, 48, 49). Immunohistochemical staining in our own laboratory has similarly failed to demonstrate dynorphin peptides in islets (data not shown), though these peptides could be detected by RIA. One possible explanation is that primary ß-cells contain insufficient quantities of dynorphin peptides for detection by immunostaining methods. This could result from rapid secretion or degradation, poor retention of peptides during tissue fixation and histologic processing, or low levels of prodynorphin gene transcription and mRNA translation. Our inability to detect prodynorphin transcripts in primary rat islets (data not shown) does not exclude possible low level expression but clearly indicates that primary cells express considerably less prodynorphin RNA than ß-cell lines. These results suggest that ß-cells may be permissive for high level prodynorphin expression only under conditions of a transformed phenotype. In this context, it is relevant to note that expression of endogenous opioids is frequently observed in primary human islet cell tumors (54).

The regulatory role of dynorphin peptides in neuronal cell function and close similarity of neurons to ß-cells (55) suggest that islet cell secretion of opioid peptides could influence ß-cell function through autocrine or paracrine effects. Islet cells of rat pancreas express µ- and {delta}-opiate-binding sites, as well as lower levels of {kappa}-receptors (15). A number of in vitro studies have demonstrated direct stimulatory effects of dynorphin peptides on cultured rodent islets. At nanomolar concentrations dynorphin-A increased glucose-induced insulin release approximately 2-fold (56, 57, 58). In contrast to its effects on neurons, dynorphin-A also increased basal islet cell cytoplasmic cAMP levels and Ca++ uptake (56, 57). At higher concentrations, effects on insulin secretion were reduced or reversed (57, 58, 59). Other opioid peptides have typically shown similar but generally weaker effects (60). Additionally, Met- and Leu-enkephalins have also been shown to increase steady state insulin RNA levels in cultured islets (16).

While potent effects of opioids on ß-cells activity have been demonstrated in vitro, examination of in vivo effects has yielded conflicting results with marked variation between species. Dynorphin-A had little effect on serum glucose or insulin in normal mice (61, 62), but significantly increased serum insulin in genetically obese (ob/ob) mice (61). Other in vivo studies in mice have shown inhibition of stimulated insulin secretion by Met-enkephalin (62). All of these studies, however, are complicated by the extremely short half-lives of these compounds in circulation (63) and potential direct effects on both {alpha}-cell glucagon secretion and hepatic glucose metabolism (64). In summary, although the presence of opioid peptides and their receptors in islets suggest potential autocrine or paracrine roles in regulating islet cell function, the significance of these mechanisms in vivo remains to be determined.

Received April 7, 1998.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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