Ascaris suum, an Intestinal Parasite, Produces Morphine1

Ascaris suum, an Intestinal Parasite, Produces Morphine This information is current as of March 24, 2015.

Yannick Goumon, Federico Casares, Stephen Pryor, Lee Ferguson, Bruce Brownawell, Patrick Cadet, Christos M. Rialas, Ingeborg D. M. Welters, Dario Sonetti and George B. Stefano J Immunol 2000; 165:339-343; ; doi: 10.4049/jimmunol.165.1.339 http://www.jimmunol.org/content/165/1/339

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The Journal of Immunology is published twice each month by The American Association of Immunologists, Inc., 9650 Rockville Pike, Bethesda, MD 20814-3994. Copyright © 2000 by The American Association of Immunologists All rights reserved. Print ISSN: 0022-1767 Online ISSN: 1550-6606.

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References

Ascaris suum, an Intestinal Parasite, Produces Morphine1 Yannick Goumon,* Federico Casares,*† Stephen Pryor,* Lee Ferguson,† Bruce Brownawell,† Patrick Cadet,* Christos M. Rialas,* Ingeborg D. M. Welters,* Dario Sonetti,‡ and George B. Stefano2*

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uccessful parasitism, in which the host survives for extended periods, can be characterized as an equilibrium between the parasite and the host, more specifically between the host’s immune system and the parasite’s ability to create a permissive microenvironment in situ. One mechanism that a parasite may use to modify the host immune response is to downregulate the host’s response (1–3). Capron and colleagues (4 –7) suggested that parasites may communicate with their hosts via common signaling molecules that diminish host immune surveillance. In this regard, morphine is generally acknowledged as an immune down-regulating agent (8). This finding is enhanced by the fact that morphine is present in several mammalian tissues, including brain and adrenal gland (9 –20), supporting its role as a neural or inflammatory mediator. Recently, we have demonstrated that free-living and parasitic invertebrates produce several major opioid peptide precursors, i.e., prodynorphin, proopiomelanocortin, and proenkephalin (21). These mammalian-like opioid peptides exhibit high sequence identity to their mammalian counterparts. For example, Mytilus adrenocorticotropin has greater than 90% sequence identity with its mammalian counterpart (21). We have also identified a tentative morphine-like molecule in Schistosoma mansoni by way of radioimmunoassay (22). Given this and the fact that the pig intestinal parasite Ascaris suum can live in its host for extended periods of time, we surmised

*Neuroscience Research Institute, State University of New York, Old Westbury, NY 11568; †Marine Sciences Research Center, State University of New York, Stony Brook, NY 11794; and ‡Neurobiology Research Unit, Department of Animal Biology, University of Modena, Modena, Italy Received for publication January 3, 2000. Accepted for publication April 12, 2000. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. 1 This work was supported by the following grants: National Institute of Mental Health (NIMH) Career Opportunities Research Program 17138; National Institute on Drug Abuse (NIDA) 09010; NIMH 47392, and the Research Foundation and Central Administration of the State University of New York and National Institutes of Health Fogarty International 00045. P.C. is a NIDA Postdoctoral Fellow. 2 Address correspondence and reprint requests to Dr. George B. Stefano, Neuroscience Research Institute, State University of New York, College at Old Westbury, Old Westbury, NY 11568. E-mail address: [email protected]

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that it might be using morphine to escape detection by the host’s immune system. In this study, we report for the first time that A. suum synthesizes morphine, thereby strengthening the commonsignal molecule hypothesis, i.e., using either similar or identical host signaling to escape host immunosurveillance.

Materials and Methods Adult frozen A. suum were obtained from Carolina Biological Supply (Burlington, NC). Live adult Ascaris were obtained from Josef Miller (Pine Plains, NY) and were maintained in the laboratory for up to 6 days as described elsewhere in detail (23). Animals were examined on the fifth day for endogenous morphine levels.

Extraction The extraction experiments using internal or external morphine standards were performed in different rooms to avoid morphine contamination of Ascaris samples. Single-use siliconized tubes also were used to prevent the loss of morphine as well as contamination between assays (14). Tissues were extensively washed with PBS buffer (three times, 1 min) to avoid an additional potential source of morphine contamination. Tissues were weighed, homogenized in 1 N HCl (1 ml/0.1 g), and then extracted with 5 ml chloroform/isopropanol 9:1 (24). After 5 min, homogenates were centrifuged (3,000 rpm, 15 min). The supernatant was centrifuged twice (15 min, 12,000 rpm, 4°C), dried, and dissolved in 0.05% trifluoroacetic acid (25). Samples were loaded on a Sep-Pak Plus cartridge (Waters, Milford, MA) and eluted in water/acetonitrile/trifluoroacetic acid (89.95%:10%:0.05%, v/v/v). Samples were dissolved in buffer A before HPLC analysis (see below).

HPLC and electrochemical detection HPLC separations were performed with two Acuflow Series VI and a C18 Unijet microbore column (Bioanalytical Systems, West Lafayette, IN). Morphine detection was performed with an amperometric detector LC-4C (Bioanalytical Systems) using a Unijet glassy carbon working electrode (3 mm) and a 0.02 Hz filter (500 mV; range, 5 nA). The mobile phases were buffer A (10 mM ammonium bicarbonate, 10 mM ammonium chloride, 10 mM sodium chloride, 0.2 mM EDTA (pH 5.0)) and buffer B (10 mM ammonium bicarbonate, 10 mM ammonium chloride, 10 mM sodium chloride, 0.2 mM EDTA, 50% acetonitile (pH 5.0) (24)). The injection volume was 5 ␮l, and the flow rate was 70 ␮l/min. Conditions were: t ⫽ 0, 0% buffer B; t ⫽ 10 min, 10% buffer B; t ⫽ 20 min, 30% buffer B; and t ⫽ 25 min, 100% buffer B. Several HPLC purifications were performed between each sample to prevent residual morphine contamination remaining on the column. Furthermore, the fraction of blank chromatography corresponding 0022-1767/00/$02.00

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The parasitic worm Ascaris suum contains the opiate alkaloid morphine as determined by HPLC coupled to electrochemical detection and by gas chromatography/mass spectrometry. The level of this material is 1168 ⴞ 278 ng/g worm wet weight. Furthermore, Ascaris maintained for 5 days contained a significant amount of morphine, as did their medium, demonstrating their ability to synthesize the opiate alkaloid. To determine whether the morphine was active, we exposed human monocytes to the material, and they immediately released nitric oxide in a naloxone-reversible manner. The anatomic distribution of morphine immunoreactivity reveals that the material is in the subcuticle layers and in the animals’ nerve chords. Furthermore, as determined by RT-PCR, Ascaris does not express the transcript of the neuronal ␮ receptor. Failure to demonstrate the expression of this opioid receptor, as well as the morphine-like tissue localization in Ascaris, suggests that the endogenous morphine is intended for secretion into the microenvironment. The Journal of Immunology, 2000, 165: 339 –343.

340 to the elution time of the morphine (flat line) was determined by gas chromatography/mass spectrometry (GC/MS)3 analysis, confirming that no morphine was remaining. Five nanograms of internal morphine standard was first chromatographed (Fig. 1A) along with Ascaris morphine (Fig. 1B), and then the same sample with 4 ng of morphine external standard was added after the extraction (Fig. 1C).

GC/MS

Bioassay of Ascaris morphine: NO release Human peripheral monocytes (Long Island Blood Services, Melville, NY) were isolated using the Accurate Scientific (Westbury, NY) monocyte kit and were washed as previously described (27–29). We used monocytes to measure the biological activity of Ascaris-derived morphine because we have demonstrated that they contain stereospecific opiate alkaloid-selective receptors that are coupled to morphine and NO release that is naloxone reversible (27, 28). NO release from the incubated monocytes (107 cells/chamber) was measured directly using an NO-specific amperometric probe (World Precision Instruments, Sarasota, FL) (30, 31). Briefly, the cells were placed in a superfusion chamber in 1 ml PBS. The probe was positioned 15 ␮m above the cell surface by a micromanipulator (Zeiss-Eppendorff, Suwanee, GA) attached to the stage of an inverted microscope (Diaphot; Nikon, Melville, NY). The system was calibrated daily by adding potassium nitrite to a solution of potassium iodide, resulting in the liberation of a known quantity of NO (World Precision Instruments). Baseline levels of NO release were determined by evaluation of NO release in PBS. Cells were stimulated with the respective purified morphine-like material, and the concentration of NO gas in solution was measured in real time with the DUO 18 computer data acquisition system (World Precision Instruments). The amperometric probe was equilibrated for at least 12 h in PBS before being transferred to the superfusion chamber containing the cells, and manipulation of the cells was performed only with glass instruments. To evaluate NO release, the cells were exposed to morphine as indicated.

Morphine immunohistochemistry Adult specimens of A. suum were obtained from Italcarni (Carpi, Italy) immediatly after the slaughter of parasitized pigs. The worms were fixed either in phosphate-buffered formalin (4% paraformaldehyde in 0.1 M (pH 7.4)) for 8 h at 4°C or in a mixture of glutaraldehyde/picric acid/acetic acid (GPA; 1:3 ⫹ 1%) overnight and then for 48 h at 4°C. Specimens were rinsed in 70% ethanol, dehydrated, embedded in paraffin, and sectioned (8 ␮m). Morphine was localized by indirect immunofluorescence using a polyclonal Ab with minimal cross-reactivity to codeine (Biogenesis, Bournemouth, U.K.). Briefly, rehydrated sections were washed in PBS (0.9% NaCl, 0.01 M sodium phosphate buffer (pH 7.4)), preincubated in 5% normal serum for 30 min, and then incubated overnight at 4°C in a moist chamber with the primary Ab. Sections were then washed with PBS 3 Abbreviations used in this paper: GC/MS, gas chromatography/mass spectrometry; MOR-IR, immunoreactive morphine.

and incubated with anti-sheep rabbit IgG-FITC (Dako, Aarhus, Denmark; 1:40; 1 h; room temperature). Slides were rinsed in PBS and then dipped in 0.005% propidium iodide for nuclei staining (1 min). After a final rapid rinse with PBS, sections were mounted in buffered glycerol (87.7% glycerol, 2.3% 1,4-diazalbicyclo [2.2.2]octane, 10% Tris 20 mM (pH 8)), cover-slipped, and examined on a Zeiss microscope equipped for fluorescence with FITC excitation and barrier filter combination. For comparison, we tested two other antisera, anti-adrenocorticotrophic hormone and anti-␤-endorphin (Biogenesis). Reaction specificity for each technique was controlled by: 1) replacing the Ab with a nonimmune serum; 2) omitting the first antiserum; and 3) preincubating the primary Ab at 4°C for 24 h in the presence of excess Ag (morphine HCl 10⫺3 M).

␮ Opiate receptor expression: isolation of total RNA Tissue samples, excised pedal ganglia of Mytilus as a positive control (32), and all body sections of Ascaris were homogenized in Tri-Reagent (Molecular Research Center, Cincinnati, OH) containing 1-bromo-3-chloropropane (0.1 ml/ml Tri-Reagent) using a Polytron homogenizer. The homogenates were stored at room temperature for 5 min to allow complete dissociation of nucleoprotein. The samples were vortexed vigorously (15 s), kept at room temperature for 7 min, and then centrifuged (15 min, 12,000 ⫻ g). The aqueous phase was transferred to a fresh tube, and the RNA was precipitated with isopropanol, washed with 75% ethanol, airdried, and resuspended in water. RNA was analyzed on a 1% denaturing agarose gel, and purity was determined spectrophotometrically.

RT-PCR of total RNA First-strand cDNA synthesis was performed using random hexamers (Life Technologies, Gaithersburg, MD). Three micrograms of total RNA isolated from pedal ganglia as positive control or Ascaris tissue was denatured at 95°C and reverse transcribed at 42°C for 1 h using Superscript II RNase H-RT (Life Technologies). Seven microliters of the reverse transcription product was added to the PCR mix containing specific primers for the ␮ opioid receptor gene and Taq DNA polymerase (Life Technologies). The PCR reaction was denatured at 95°C for 5 min before 30 cycles at 95°C for 1 min, 57°C for 1 min, 72°C for 1 min, and then an extension step cycle at 72°C for 10 min. PCR products were analyzed on a 2% agarose gel (Sigma, St. Louis, MO) and visualized by ethidium bromide staining. The ␮-specific primers used in the PCR reactions amplified a 441-bp fragment starting at map position 896 (primer M1, 5⬘-GGTACTGG GAAAACCTGCTGAAGATCTGTG-3⬘) and at map position 1336 (primer M4, 5⬘-GGTCTCTAGTGTTCTGACGAATTCGAGTGG-3⬘). This segment of the gene encodes the third extracellular loop of the receptor that is important for ␮ agonist selectivity. In addition, primers for the internal control gene G3PDH (forward primer, 5⬘-ACCACAGTCCATGCCAT CAC; reverse primer, 5⬘-TCCACCACCCTGTTGCTGGTA) were used to amplify a 451-bp fragment by RT-PCR from Ascaris total RNA.

Results To determine whether the parasitic worm A. suum utilized the opioid alkaloid morphine, this compound was identified and quantified, and then the worms were analyzed for the presence of morphine receptors. Morphine was identified in Ascaris extracts by reversed-phase HPLC using a gradient of acetonitrile after liquid and solid extraction (Fig. 1). All experiments were carefully performed to prevent exogenous morphine contamination (see Materials and Methods). The morphine extracted from Ascaris (Fig. 1B) was identical with the major peak for the morphine internal (Fig. 1A) and external (Fig. 1C) standard. This finding was replicated in five other worms. The linearity of the detection of the HPLC technique at low concentrations of morphine substantiates the sensitivity of this method for determining morphine levels (Fig. 1B, inset). The concentration of morphine was determined using the Chromatogram Report (Bioanalytical Systems) and extrapolated from the peak area calculated for the internal standard. The average concentration of morphine in the six samples was 1168 ⫾ 278 ng/g wet weight, and after 5 days in culture it was 41 ⫾ 15 ng/g wet weight. In the worm incubation medium (nine adult worms in 1.48 L) changed daily, on the fifth day morphine also was present at 725

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The identity of the morphine detected by HPLC was further confirmed by GC/MS using a modification of a method reported by Allen et al. (26). A fraction of the eluent corresponding to the HPLC morphine peak was collected in a 0.5-ml siliconized plastic vial, evaporated to dryness, and stored at ⫺70°C. The sample was dissolved in 50 ␮l of 2% ammonia (in methanol) and transferred to 0.1-ml conical glass vials (Kimax, Alphuretta, GA) fitted with Teflon-lined septum liners. The sample was evaporated under nitrogen gas and then mixed with N,O-bis (trimethylsilyl) trifluoroacetimide (20 –50 ␮l) catalyzed with 1% trimethylchlorosilane (90°C, 30 min). This reaction yields a ditrimethylsilyl derivative of morphine (m.w. ⫽ 429). The reaction conditions chosen resulted in higher yields than did reactions conducted at lower temperature (70°C) or for shorter times. The derivatized morphine was analyzed on a Varian Saturn III GC/MS equipped with a 30-m (0.25 mm internal diameter and 0.25 ␮m) DB-5 capillary column (J & W Scientific, Folsom, CA). Two-microliter injections were made in splitless mode at an initial column temperature of 180°C, and after 2 min the column temperature was heated at a rate of 15°C/min up to 300°C. Morphine was detected at 10 min; injection port temperature was 275°C. Mass spectrum indicated major ions (base peak depended on instrument tune conditions) at 429 (M⫹) and 414 (M-CH3⫹). Analyses of samples were conducted using a selected ion storage method, in which mass windows (⫾ 2 atomic mass units) around ions 429 and 414 were collected. Morphine identity was confirmed by the retention time, peak shape, and comparison of parasite-derived morphine to authentic morphine standards injected.

PARASITE MAKES MORPHINE

The Journal of Immunology

341 nolocalization (data not shown). There was no staining in sections in which the primary antiserum was substituted with normal serum. We next determined whether tissues from Ascaris have opiate receptors that would allow them to utilize the morphine they contain. We used RT-PCR to amplify a fragment of the coding region of the ␮ opiate receptor from Ascaris and also one from Mytilus as a positive control (Fig. 3). Using ␮-specific primers, we isolated a transcript of the expected size for the ␮ receptor (441 bp) from Mytilus as positive contol (Ref. 32 and Fig. 3, lane 2) but not from Ascaris (Fig. 3, lane 3). This was not due to a lack of mRNA from Ascaris, because we were able to amplify a 451-bp mRNA corresponding to G3PDH from the worm (34). Sequence analysis of the Mytilus PCR product demonstrated that the ␮ receptor fragment exhibited 95% sequence identity with the human brain ␮ opiate receptor (32). Failure to demonstrate the ␮ receptor in Ascaris tissues suggests that this material is intended for secretion into the microenvironment as suggested by the MOR-IR localization.

FIGURE 1. Purification of morphine from Ascaris. Morphine was isolated by HPLC. Running conditions: range, 5 nA; filter, 0.02 Hz; potential, 500 mV; flow rate, 500 ␮l/min; A buffer: 1 mM ammonium bicarbonate, 10 mM ammonium chloride, 10 mM sodium chloride, 0.2 mM EDTA (pH 5.0); B buffer: same as A buffer but with 50% acetonitrile. Gradient: t ⫽ 0, 0% buffer B; t ⫽ 10 min, 10% buffer B; t ⫽ 20 min, 30% buffer B; t ⫽ 25 min, 100% buffer B. A, Five nanograms of morphine internal standard. B, Standard curve of low morphine concentrations. C, Ascaris extract. D, Ascaris extract ⫹ 4 ng morphine external standard. E, Ascaris incubation medium.

ng/L, demonstrating that the worm was synthesizing and secreting the material (Fig. 1E). The morphine in the HPLC fractions was further analyzed by GC/MS (Fig. 2A). Again, the morphine fraction isolated from A. suum was identical with synthetic morphine according to retention time, peak shape, and comparison to authentic morphine. Morphine quantification was confirmed by this method. Additionally, the identity of morphine in the worm incubation medium, noted above, was confirmed by GC/MS (data not shown). To determine whether the morphine was active, we used a classic bioassay for morphine, i.e., the ability of the compound of interest to release NO from human monocytes in a naloxone-reversible manner (28). Incubation of human monocytes with morphine from Ascaris resulted in the immediate release of NO that was antagonized by naloxone (Fig. 2B). We next analyzed the anatomic distribution of morphine in A. suum by immunofluorescence using a morphine-specific Ab (Ref. 33 and Fig. 2, C and D). Immunoreactive morphine (MOR-IR; green fluorescence) was found in subcuticle layers among collagenous structures, in fiber-like structures in the hypodermis, in the radial intercellular spaces between longitudinal contracting muscles underlying the skin, in the spaces encircling the inner big bodies of muscle cells, and in the nerve chords. Additionally, the animal’s ova appear to be surrounded by this material in the extracellular space. Different fixation methods gave the same results. Staining for adrenocorticotrophic hormone and ␤-endorphin were negative, further confirming the specificity of the morphine immu-

The present study demonstrates for the first time the presence of morphine in A. suum and its expression under presumed nonstimulated (or basal) conditions. Animals maintained in the laboratory for 5 days also had morphine present and were secreting the material into the medium, demonstrating their ability to synthesize this material. Unlike the free-living mollusk Mytilus edulis (32), Ascaris does not express ␮ opiate receptor transcripts. In Ascaris, MOR-IR was found in the epidermis of the animal. These two pieces of data suggest that, in Ascaris, morphine is secreted into the microenvironment where it could be used as a signaling molecule. We hypothesize that the morphine helps the parasite to evade the immunosurveillance machinery of the host and thereby increase the stability of its microenvironment and ensure its survival. This strategy appears to also ensure the survival of the ova. Furthermore, the morphine Ascaris displays the same effect as natural morphine on the NO release of the monocyte, an effect antagonized by the naloxone. This result suggests that the morphine from Ascaris can act on immune cells to enhance its survival. Opioid peptides and their precursors are present in free-living and parasitic invertebrates (21). These free-living invertebrates contain complex and highly specific opioid and opiate receptor sites (32, 35–37) involved in dopaminergic and NO processes (31, 38). In these animals, the tissues that produced these signaling molecules have the appropriate receptors to utilize them. For example, in Mytilus, we recently demonstrated both morphine and a ␮ opiate receptor subtype that exhibited 95% sequence identity with the human neuronal ␮ opiate receptor (32). We had surmised, based on the Mytilus model, that the signaling material was used within that tissue. However, Ascaris makes morphine without a corresponding receptor, suggesting that it is secreted for “external” signaling, i.e., host immune down-regulation (8, 39). This hypothesis is supported by our immunohistochemistry results, which demonstrated the presence of this material in the epidermis of the animal. Complementing the present study are reports that Schistosoma synthesize a morphine-like compound inhibiting immunocytes in a naloxone-reversible manner (22). In this report, we also demonstrated that in worms maintained in the laboratory the endogenous morphine-like levels were low and that, upon exposure to human immunocytes, a significant increase in the worm’s opiate level occurred, suggesting a feedback process. In the present report we also note a significant drop in morphine levels in animals maintained in the laboratory. However, its high level in the incubation

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Discussion

342

PARASITE MAKES MORPHINE

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FIGURE 2. Characterization of Ascaris morphine. A, GC/MS spectrum of two morphine standards (200 pg and 400 pg) and the morphine material collected during HPLC analysis of Ascaris. B, Bioassay of morphine-like activity extracted from Ascaris. Ascaris extracts stimulate NO release from human monocytes. Top, A total of 500 ␮l of Ascaris whole body extract was added to a solution containing purified human monocytes (107 cells/ml) (arrowhead). This resulted in NO release. Bottom, Naloxone (first arrowhead; 10⫺6 M) was added to another batch of cells under similar conditions before the addition of morphine extract (second arrowhead). This treatment blocked the Ascaris morphine-stimulated NO release, demonstrating that the Ascaris morphine material is opiate alkaloid in character. C and D, Immunocytochemical localization of morphine in Ascaris. C, Ascaris body wall region incubated with nonimmune serum shows a complete lack of positive immunoreactivity. D, MOR-IR in Ascaris suum. Transverse section of the body wall. Cuticule (a) MOR-IR is localized in the fibrous layer under the cuticle (b), in fibers running in the hypodermis (c) and in the intercellualr spaces between the contractile parts of myoepithelial cells (d) (bar ⫽ 20 microns). E, Ascaris ova incubated with nonimmune serum shows a complete lack of positive immunoreactivity. F, Morphine-like immunoreactivity immediately outside the ova (bar ⫽ 50 microns).

medium indicates that as this material is made it is secreted into the immediate environment. In this regard, if morphine were not synthesized in the worm, by day 5 it should not be detected because its tissue half-life is about 2 h and it cannot be detected in mammalian and invertebrate tissues after 24 h (Ref. 40 and our unpublished observations).

These Ascaris morphine data also suggest that morphine may play a role in gastrointestinal regulation. In human history it has widely been accepted that diarrhea represents the body’s attempt to flush out toxins, including parasitic worms. The Persian physician Avicenna prescribed morphine for cough, anemia, and diarrhea thousands of years ago (41). Thus, Ascaris may secrete this opiate

The Journal of Immunology

FIGURE 3. Identification of ␮ opioid receptor mRNA transcript. Total RNA isolated from these tissues was reverse transcribed, PCR amplified, and run on agarose gels. Lane 1, G3PDH housekeeping gene (expected transcript size is 451 bp). Lane 2, ␮ opioid receptor transcript in Mytilus pedal ganglia (expected size, 441 bp). Lane 3, lack of ␮ receptor transcript in Ascaris. Lane 4, 100-bp DNA markers. Both negative and positive strands were sequenced; however, only the sequence from the negative strand is shown in lane 2. The tissues were washed extensively with PBS to limit any contamination.

Acknowledgments We thank Danielle Benz, Ireen Khan, Mazen Madhoun (National Institute of Mental Health-Career Opportunities Research Program Fellows), and Yun Su Lee for technical assistance.

References 1. Capron, A., J. P. Dessaint, M. Capron, J. H. Ouma, and A. E. Butterworth. 1987. Immunity towards schistosomes: progress toward vaccine. Science 238:1065. 2. Capron, A., and J. P. Dessaint. 1988. Parasitism, a model of cell sociology. News Physiol. Sci. 3:75. 3. Capron, A., and J. P. Dessaint. 1989. Molecular basis of host-parasite relationship: toward the definition of protective antigens. Immunol. Rev. 112:27. 4. Duvaux-Miret, O., C. Dissous, J. P. Guatron, E. Pattou, C. Kordon, and A. Capron. 1990. The helminth Schistosoma mansoni expresses a peptide similar to human ␤-endorphin and possesses a POMC-related gene. New Biol. 2:93. 5. Duvaux-Miret, O., G. B. Stefano, E. M. Smith, C. Dissous, and A. Capron. 1992. Immunosuppression in the definitive and intermediate hosts of the human parasite Schistosoma mansoni by release of immunoactive neuropeptides. Proc. Natl. Acad. Sci. USA 89:778. 6. Duvaux-Miret, O., G. B. Stefano, E. M. Smith, L. Mallozzi, and A. Capron. 1992. Proopiomelanocortin-derived peptides as tools of immune evasion for the human trematode Schistosoma mansoni. Acta Biol. Hung. 43:281. 7. Duvaux-Miret, O., G. B. Stefano, E. M. Smith, and A. Capron. 1992. Neuroimmunology of host parasite interactions: proopiomelanocortin derived peptides in the infection by Schistosoma mansoni. Adv. Neuroimmunol. 2:297. 8. Stefano, G. B., B. Scharrer, E. M. Smith, T. K. Hughes, H. I. Magazine, T. V. Bilfinger, A. Hartman, G. L. Fricchione, Y. Liu, and M. H. Makman. 1996. Opioid and opiate immunoregulatory processes. Crit. Rev. Immunol. 16:109. 9. Donnerer, J., G. Cardinale, J. Coffey, C. A. Lisek, I. Jardine, and S. Spector. 1987. Chemical characterization and regulation of endogenous morphine and codeine in the rat. J. Pharmacol. Exp. Ther. 242:583. 10. Donnerer, J., K. Oka, A. Brossi, K. C. Rice, and S. Spector. 1986. Presence and formation of codeine and morphine in the rat. Proc. Natl. Acad. Sci. USA 83: 4566. 11. Cardinale, G. J., J. Donnerer, A. D. Finck, J. D. Kantrowitz, K. Oka, and S. Spector. 1987. Morphine and codeine are endogenous components of human cerebrospinal fluid. Life Sci. 40:301. 12. Donnerer, J., G. Cardinale, J. Coffey, C. A. Lisek, I. Jardine, and S. Spector. 1987. Chemical characterization and regulation of endogenous morphine and codeine in the rat. J. Pharmacol. Exp. Ther. 242:583. 13. Goldstein, A., R. W. Barrett, I. F. James, L. I. Lowney, C. Weitz, L. I. Knipmeyer, and H. Rapoport. 1985. Morphine and other opiates from beef brain and adrenal. Proc. Natl. Acad. Sci. USA 82:5203. 14. Weitz, C. J., L. I. Lowney, K. F. Faull, G. Feister, and A. Goldstein. 1986. Morphine and codeine from mammalian brain. Proc. Natl. Acad. Sci. USA 83: 9784. 15. Weitz, C. J., K. F. Faull, and A. Goldstein. 1987. Synthesis of the skeleton of morphine molecule by mammalian liver. Nature 330:674.

16. Weitz, C. J., L. I. Lowney, K. F. Faull, G. Feistner, and A. Goldstein. 1986. Morphine and codeine from mammalian brain. Proc. Natl. Acad. Sci. USA 83: 9784. 17. Bianchi, E., C. Alessandrini, M. Guarna, and A. Tagliamonte. 1993. Endogenous codeine and morphine are stored in specific brain neurons. Brain Res. 627:210. 18. Bianchi, E., M. Guarna, and A. Tagliamonte. 1994. Immunocytochemical localization of endogenous codeine and morphine. Adv. Neuroimmunol. 4:83. 19. Guarna, M., C. Neri, F. Petrioli, and E. Bianchi. 1998. Potassium-induced release of endogenous morphine form rat brain slices. J. Neurochem. 70:147. 20. Tagliamonte, A., M. Guarna, and E. Bianchi. 1995. Endogenous morphine and codeine as possible physiological ligands of ␮-opiate receptors. In Drug Addiction and Related Clinical Problems. I. Maremmani and A. Tagliamonte, eds. Springer-Verlag, Vienna, p. 27. 21. Stefano, G. B., and M. Salzet. 1999. Invertebrate opioid precursors: evolutionary conservation and the significance of enzymatic processing. Int. Rev. Cytol. 187: 261. 22. Leung, M. K., C. Dissous, A. Capron, H. Woldegaber, O. Duvaux-Miret, S. C. Pryor, and G. B. Stefano. 1995. Schistosoma mansoni: the presence and potential use of opiate-like substances. Exp. Parasitol. 81:208. 23. Bowman, J. W., C. A. Winterrowd, A. R. Friedman, D. P. Thompson, R. D. Klein, J. P. Davis, A. G. Maule, K. L. Blair, and T. G. Geary. 1995. Nitric oxide mediates the inhibitory effects of SDPNFLRFamide, a nematode FMRFamide-related neuropeptide, in Ascaris suum. J. Neurophysiol. 74:1880. 24. Grieco, D., A. Porcellini, E. V. Avvedimento, and M. E. Gottesman. 1996. Requirement for cAMP-PKA pathway activation by M phase-promoting factor in the transition from mitosis to interphase. Science 271:1718. 25. Macarthur, H., M. B. Mattammal, and T. C. Westfall. 1995. A new perspective on the inhibitory role of nitric oxide in sympathetic neurotransmission. Biochem. Biophys. Res. Commun. 216:686. 26. Allen, D. L., K. S. Scott, and J. S. Oliver. 1999. Comparison of solid-phase extraction and supercritical fluid extraction for the analysis of morphine in whole blood. J. Anal. Toxicol. 23:216. 27. Stefano, G. B., A. Digenis, S. Spector, M. K. Leung, T. V. Bilfinger, M. H. Makman, B. Scharrer, and N. N. Abumrad. 1993. Opiatelike substances in an invertebrate, a novel opiate receptor on invertebrate and human immunocytes, and a role in immunosuppression. Proc. Natl. Acad. Sci. USA 90:11099. 28. Magazine, H. I., Y. Liu, T. V. Bilfinger, G. L. Fricchione, and G. B. Stefano. 1996. Morphine-induced conformational changes in human monocytes, granulocytes, and endothelial cells and in invertebrate immunocytes and microglia are mediated by nitric oxide. J. Immunol. 156:4845. 29. Stefano, G. B., P. Melchiorri, L. Negri, T. K. Hughes, and B. Scharrer. 1992. (D-Ala2)-Deltorphin I binding and pharmacological evidence for a special subtype of ␦ opioid receptor on human and invertebrate immune cells. Proc. Natl. Acad. Sci. USA 89:9316. 30. Stefano, G. B., A. Hartman, T. V. Bilfinger, H. I. Magazine, Y. Liu, F. Casares, and M. S. Goligorsky. 1995. Presence of the ␮3 opiate receptor in endothelial cells: coupling to nitric oxide production and vasodilation. J. Biol. Chem. 270: 30290. 31. Liu, Y., D. Shenouda, T. V. Bilfinger, M. L. Stefano, H. I. Magazine, and G. B. Stefano. 1996. Morphine stimulates nitric oxide release from invertebrate microglia. Brain Res. 722:125. 32. Cadet, P., and G. B. Stefano. 1999. Mytilus edulis pedal ganglia express ␮ opiate receptor transcripts exhibiting high sequence identity with human neuronal ␮1. Mol. Brain Res. 74:242. 33. Sonetti, D., L. Mola, F. Casares, E. Bianchi, M. Guarna, and G. B. Stefano. 1999. Endogenous morphine levels increase in molluscan neural and immune tissues after physical trauma. Brain Res. 835:137. 34. Kochman, M., J. Golebiowska, T. Baranowski, J. R. Dedman, D. W. Fodge, and B. G. Harris. 1975. Studies on enzymes from parasitic helminths. V. Purification and characterization of glyceraldehyde 3-phosphate dehydrogenase from Ascaris suum muscle. Comp. Biochem. Physiol. 52:301. 35. Kream, R. M., R. S. Zukin, and G. B. Stefano. 1980. Demonstration of two classes of opiate binding sites in the nervous tissue of the marine mollusc Mytilus edulis: positive homotropic cooperativity of lower affinity binding sites. J. Biol. Chem. 255:9218. 36. Stefano, G. B., R. M. Kream, and R. S. Zukin. 1980. Demonstration of stereospecific opiate binding in the nervous tissue of the marine mollusc Mytilus edulis. Brain Res. 181:445. 37. Stefano, G. B., and B. Scharrer. 1981. High affinity binding of an enkephalin analog in the cerebral ganglion of the insect Leucophaea maderae (Blattaria). Brain Res. 225:107. 38. Stefano, G. B., and B. Scharrer. 1996. The presence of the ␮3 opiate receptor in invertebrate neural tissues. Comp. Biochem. Physiol. 113C:369. 39. Eisenstein, T. K., and M. E. Hilburger. 1998. Opioid modulation of immune responses: effects on phagocyte and lymphoid cell populations. J. Neuroimmunol. 83:36. 40. Reisine, T., and G. W. Pasternak. 1996. Opioid analgesics and antagonists. In The Pharmacological Basis of Therapeutics. J. G. Hardman and L. E. Limbird, eds. McGraw–Hill, New York, p. 521. 41. Merlin, M. D. 1984. On The Trail of Ancient Opium Poppy. Associated University Press, London.

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alkaloid to diminish its “flushing” from the host because a secondary effect on gastrointestinal motility is to induce constipation. In conclusion, opioid processes appear to have evolved much earlier than previously thought. Opiate alkaloid signaling appears to have been conserved during evolution and used to enhance the longevity of the host and parasite. Clearly, this represents an important strategy for enhancing the odds for survival and, thus, for simultaneously preserving the message.

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