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Morphine, an Intestinal Parasite, ProducesAscaris suum

StefanoRialas, Ingeborg D. M. Welters, Dario Sonetti and George B.Ferguson, Bruce Brownawell, Patrick Cadet, Christos M. Yannick Goumon, Federico Casares, Stephen Pryor, Lee

http://www.jimmunol.org/content/165/1/339doi: 10.4049/jimmunol.165.1.339

2000; 165:339-343; ;J Immunol 

Referenceshttp://www.jimmunol.org/content/165/1/339.full#ref-list-1

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Print ISSN: 0022-1767 Online ISSN: 1550-6606. Immunologists All rights reserved.Copyright © 2000 by The American Association of1451 Rockville Pike, Suite 650, Rockville, MD 20852The American Association of Immunologists, Inc.,

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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,‡ andGeorge B. Stefano2*

The parasitic worm Ascaris suumcontains the opiate alkaloid morphine as determined by HPLC coupled to electrochemicaldetection and by gas chromatography/mass spectrometry. The level of this material is 11686 278 ng/g worm wet weight. Fur-thermore, Ascarismaintained for 5 days contained a significant amount of morphine, as did their medium, demonstrating theirability to synthesize the opiate alkaloid. To determine whether the morphine was active, we exposed human monocytes to thematerial, and they immediately released nitric oxide in a naloxone-reversible manner. The anatomic distribution of morphineimmunoreactivity reveals that the material is in the subcuticle layers and in the animals’ nerve chords. Furthermore, as deter-mined by RT-PCR, Ascarisdoes not express the transcript of the neuronalm receptor. Failure to demonstrate the expression ofthis opioid receptor, as well as the morphine-like tissue localization inAscaris, suggests that the endogenous morphine is intendedfor secretion into the microenvironment. The Journal of Immunology,2000, 165: 339–343.

Successful parasitism, in which the host survives for ex-tended periods, can be characterized as an equilibrium be-tween the parasite and the host, more specifically between

the host’s immune system and the parasite’s ability to create apermissive microenvironment in situ. One mechanism that a par-asite may use to modify the host immune response is to down-regulate the host’s response (1–3). Capron and colleagues (4–7)suggested that parasites may communicate with their hosts viacommon signaling molecules that diminish host immune surveil-lance. In this regard, morphine is generally acknowledged as animmune down-regulating agent (8). This finding is enhanced bythe fact that morphine is present in several mammalian tissues,including brain and adrenal gland (9–20), supporting its role as aneural or inflammatory mediator.

Recently, we have demonstrated that free-living and parasiticinvertebrates produce several major opioid peptide precursors, i.e.,prodynorphin, proopiomelanocortin, and proenkephalin (21).These mammalian-like opioid peptides exhibit high sequence iden-tity to their mammalian counterparts. For example,Mytilus adre-nocorticotropin has greater than 90% sequence identity with itsmammalian counterpart (21). We have also identified a tentativemorphine-like molecule inSchistosoma mansoniby way of radio-immunoassay (22).

Given this and the fact that the pig intestinal parasiteAscarissuumcan live in its host for extended periods of time, we surmised

that it might be using morphine to escape detection by the host’simmune system. In this study, we report for the first time thatA.suumsynthesizes morphine, thereby strengthening the common-signal molecule hypothesis, i.e., using either similar or identicalhost signaling to escape host immunosurveillance.

Materials and MethodsAdult frozenA. suumwere obtained from Carolina Biological Supply (Bur-lington, NC). Live adultAscariswere obtained from Josef Miller (PinePlains, NY) and were maintained in the laboratory for up to 6 days asdescribed elsewhere in detail (23). Animals were examined on the fifth dayfor endogenous morphine levels.

Extraction

The extraction experiments using internal or external morphine standardswere performed in different rooms to avoid morphine contamination ofAscarissamples. Single-use siliconized tubes also were used to prevent theloss of morphine as well as contamination between assays (14). Tissueswere extensively washed with PBS buffer (three times, 1 min) to avoid anadditional potential source of morphine contamination. Tissues wereweighed, homogenized in 1 N HCl (1 ml/0.1 g), and then extracted with 5ml chloroform/isopropanol 9:1 (24). After 5 min, homogenates were cen-trifuged (3,000 rpm, 15 min). The supernatant was centrifuged twice (15min, 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) andeluted 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 C18Unijet 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 (3mm) and a 0.02 Hz filter (500 mV; range, 5 nA). The mobile phases werebuffer A (10 mM ammonium bicarbonate, 10 mM ammonium chloride, 10mM sodium chloride, 0.2 mM EDTA (pH 5.0)) and buffer B (10 mMammonium bicarbonate, 10 mM ammonium chloride, 10 mM sodium chlo-ride, 0.2 mM EDTA, 50% acetonitile (pH 5.0) (24)). The injection volumewas 5ml, and the flow rate was 70ml/min. Conditions were:t 5 0, 0%buffer B; t 5 10 min, 10% buffer B;t 5 20 min, 30% buffer B; andt 5 25min, 100% buffer B. Several HPLC purifications were performed betweeneach sample to prevent residual morphine contamination remaining on thecolumn. Furthermore, the fraction of blank chromatography corresponding

*Neuroscience Research Institute, State University of New York, Old Westbury, NY11568; †Marine Sciences Research Center, State University of New York, StonyBrook, 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 pagecharges. This article must therefore be hereby markedadvertisementin accordancewith 18 U.S.C. Section 1734 solely to indicate this fact.1 This work was supported by the following grants: National Institute of MentalHealth (NIMH) Career Opportunities Research Program 17138; National Institute onDrug Abuse (NIDA) 09010; NIMH 47392, and the Research Foundation and CentralAdministration of the State University of New York and National Institutes of HealthFogarty International 00045. P.C. is a NIDA Postdoctoral Fellow.2 Address correspondence and reprint requests to Dr. George B. Stefano, Neuro-science Research Institute, State University of New York, College at Old Westbury,Old Westbury, NY 11568. E-mail address: GBS11@banet.net

Copyright © 2000 by The American Association of Immunologists 0022-1767/00/$02.00

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to the elution time of the morphine (flat line) was determined by gas chro-matography/mass spectrometry (GC/MS)3 analysis, confirming that nomorphine was remaining. Five nanograms of internal morphine standardwas first chromatographed (Fig. 1A) along withAscaris morphine (Fig.1B), and then the same sample with 4 ng of morphine external standard wasadded after the extraction (Fig. 1C).

GC/MS

The identity of the morphine detected by HPLC was further confirmed byGC/MS using a modification of a method reported by Allen et al. (26). Afraction of the eluent corresponding to the HPLC morphine peak was col-lected in a 0.5-ml siliconized plastic vial, evaporated to dryness, and storedat 270°C. The sample was dissolved in 50ml of 2% ammonia (in meth-anol) and transferred to 0.1-ml conical glass vials (Kimax, Alphuretta, GA)fitted with Teflon-lined septum liners. The sample was evaporated undernitrogen gas and then mixed withN,O-bis (trimethylsilyl) trifluoroacetim-ide (20–50ml) catalyzed with 1% trimethylchlorosilane (90°C, 30 min).This reaction yields a ditrimethylsilyl derivative of morphine (m.w.5429). The reaction conditions chosen resulted in higher yields than didreactions conducted at lower temperature (70°C) or for shorter times. Thederivatized morphine was analyzed on a Varian Saturn III GC/MSequipped with a 30-m (0.25 mm internal diameter and 0.25mm) DB-5capillary column (J & W Scientific, Folsom, CA). Two-microliter injec-tions were made in splitless mode at an initial column temperature of180°C, and after 2 min the column temperature was heated at a rate of15°C/min up to 300°C. Morphine was detected at 10 min; injection porttemperature was 275°C. Mass spectrum indicated major ions (base peakdepended on instrument tune conditions) at 429 (M1) and 414 (M-CH31).Analyses of samples were conducted using a selected ion storage method,in which mass windows (62 atomic mass units) around ions 429 and 414were collected. Morphine identity was confirmed by the retention time,peak shape, and comparison of parasite-derived morphine to authentic mor-phine standards injected.

Bioassay ofAscarismorphine: NO release

Human peripheral monocytes (Long Island Blood Services, Melville, NY)were isolated using the Accurate Scientific (Westbury, NY) monocyte kitand were washed as previously described (27–29). We used monocytes tomeasure the biological activity ofAscaris-derived morphine because wehave demonstrated that they contain stereospecific opiate alkaloid-selectivereceptors that are coupled to morphine and NO release that is naloxonereversible (27, 28).

NO release from the incubated monocytes (107 cells/chamber) was mea-sured directly using an NO-specific amperometric probe (World PrecisionInstruments, Sarasota, FL) (30, 31). Briefly, the cells were placed in asuperfusion chamber in 1 ml PBS. The probe was positioned 15mm abovethe 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 asolution of potassium iodide, resulting in the liberation of a known quantityof NO (World Precision Instruments). Baseline levels of NO release weredetermined by evaluation of NO release in PBS. Cells were stimulated withthe respective purified morphine-like material, and the concentration of NOgas in solution was measured in real time with the DUO 18 computer dataacquisition system (World Precision Instruments). The amperometric probewas equilibrated for at least 12 h in PBS before being transferred to thesuperfusion chamber containing the cells, and manipulation of the cells wasperformed only with glass instruments. To evaluate NO release, the cellswere exposed to morphine as indicated.

Morphine immunohistochemistry

Adult specimens ofA. suumwere obtained from Italcarni (Carpi, Italy)immediatly after the slaughter of parasitized pigs. The worms were fixedeither in phosphate-buffered formalin (4% paraformaldehyde in 0.1 M (pH7.4)) for 8 h at 4°C or in a mixture of glutaraldehyde/picric acid/acetic acid(GPA; 1:3 1 1%) overnight and then for 48 h at 4°C. Specimens wererinsed in 70% ethanol, dehydrated, embedded in paraffin, and sectioned (8mm). Morphine was localized by indirect immunofluorescence using apolyclonal 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 in5% normal serum for 30 min, and then incubated overnight at 4°C in amoist chamber with the primary Ab. Sections were then washed with PBS

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 iodidefor nuclei staining (1 min). After a final rapid rinse with PBS, sections weremounted 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 aZeiss microscope equipped for fluorescence with FITC excitation and bar-rier filter combination.

For comparison, we tested two other antisera, anti-adrenocorticotrophichormone and anti-b-endorphin (Biogenesis). Reaction specificity for eachtechnique was controlled by: 1) replacing the Ab with a nonimmune serum;2) omitting the first antiserum; and 3) preincubating the primary Ab at 4°Cfor 24 h in the presence of excess Ag (morphine HCl 1023 M).

m Opiate receptor expression: isolation of total RNA

Tissue samples, excised pedal ganglia ofMytilus as a positive control (32),and all body sections ofAscariswere homogenized in Tri-Reagent (Mo-lecular Research Center, Cincinnati, OH) containing 1-bromo-3-chloropro-pane (0.1 ml/ml Tri-Reagent) using a Polytron homogenizer. The homog-enates were stored at room temperature for 5 min to allow completedissociation of nucleoprotein. The samples were vortexed vigorously (15s), kept at room temperature for 7 min, and then centrifuged (15 min,12,0003 g). The aqueous phase was transferred to a fresh tube, and theRNA was precipitated with isopropanol, washed with 75% ethanol, air-dried, and resuspended in water. RNA was analyzed on a 1% denaturingagarose gel, and purity was determined spectrophotometrically.

RT-PCR of total RNA

First-strand cDNA synthesis was performed using random hexamers (LifeTechnologies, Gaithersburg, MD). Three micrograms of total RNA isolatedfrom pedal ganglia as positive control orAscaristissue was denatured at95°C and reverse transcribed at 42°C for 1 h using Superscript II RNaseH-RT (Life Technologies). Seven microliters of the reverse transcriptionproduct was added to the PCR mix containing specific primers for themopioid receptor gene andTaq DNA polymerase (Life Technologies). ThePCR reaction was denatured at 95°C for 5 min before 30 cycles at 95°C for1 min, 57°C for 1 min, 72°C for 1 min, and then an extension step cycleat 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 m-specific primers used in the PCR reactions amplified a 441-bpfragment starting at map position 896 (primer M1, 59-GGTACTGGGAAAACCTGCTGAAGATCTGTG-39) and at map position 1336 (prim-er M4, 59-GGTCTCTAGTGTTCTGACGAATTCGAGTGG-39). This seg-ment of the gene encodes the third extracellular loop of the receptor that isimportant for m agonist selectivity. In addition, primers for the internalcontrol gene G3PDH (forward primer, 59-ACCACAGTCCATGCCATCAC; reverse primer, 59-TCCACCACCCTGTTGCTGGTA) were used toamplify a 451-bp fragment by RT-PCR fromAscaristotal RNA.

ResultsTo determine whether the parasitic wormA. suumutilized the opi-oid alkaloid morphine, this compound was identified and quanti-fied, and then the worms were analyzed for the presence of mor-phine receptors. Morphine was identified inAscaris extracts byreversed-phase HPLC using a gradient of acetonitrile after liquidand solid extraction (Fig. 1). All experiments were carefully per-formed to prevent exogenous morphine contamination (seeMate-rials and Methods).The morphine extracted fromAscaris (Fig.1B) was identical with the major peak for the morphine internal(Fig. 1A) and external (Fig. 1C) standard. This finding was repli-cated in five other worms. The linearity of the detection of theHPLC technique at low concentrations of morphine substantiatesthe sensitivity of this method for determining morphine levels (Fig.1B, inset).

The concentration of morphine was determined using the Chro-matogram Report (Bioanalytical Systems) and extrapolated fromthe peak area calculated for the internal standard. The averageconcentration of morphine in the six samples was 11686 278 ng/gwet weight, and after 5 days in culture it was 416 15 ng/g wetweight. In the worm incubation medium (nine adult worms in 1.48L) changed daily, on the fifth day morphine also was present at 725

3 Abbreviations used in this paper: GC/MS, gas chromatography/mass spectrometry;MOR-IR, immunoreactive morphine.

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ng/L, demonstrating that the worm was synthesizing and secretingthe material (Fig. 1E).

The morphine in the HPLC fractions was further analyzed byGC/MS (Fig. 2A). Again, the morphine fraction isolated fromA.suumwas identical with synthetic morphine according to retentiontime, peak shape, and comparison to authentic morphine. Mor-phine quantification was confirmed by this method. Additionally,the identity of morphine in the worm incubation medium, notedabove, was confirmed by GC/MS (data not shown).

To determine whether the morphine was active, we used a clas-sic bioassay for morphine, i.e., the ability of the compound ofinterest to release NO from human monocytes in a naloxone-re-versible manner (28). Incubation of human monocytes with mor-phine fromAscarisresulted in the immediate release of NO thatwas antagonized by naloxone (Fig. 2B).

We next analyzed the anatomic distribution of morphine inA.suumby 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 collag-enous structures, in fiber-like structures in the hypodermis, in theradial intercellular spaces between longitudinal contracting mus-cles underlying the skin, in the spaces encircling the inner bigbodies of muscle cells, and in the nerve chords. Additionally, theanimal’s ova appear to be surrounded by this material in the ex-tracellular space. Different fixation methods gave the same results.Staining for adrenocorticotrophic hormone andb-endorphin werenegative, further confirming the specificity of the morphine immu-

nolocalization (data not shown). There was no staining in sectionsin which the primary antiserum was substituted with normalserum.

We next determined whether tissues fromAscarishave opiatereceptors that would allow them to utilize the morphine they con-tain. We used RT-PCR to amplify a fragment of the coding regionof them opiate receptor fromAscarisand also one fromMytilus asa positive control (Fig. 3). Usingm-specific primers, we isolated atranscript of the expected size for them receptor (441 bp) fromMytilus as positive contol (Ref. 32 and Fig. 3,lane 2) but notfrom Ascaris(Fig. 3, lane 3). This was not due to a lack of mRNAfrom Ascaris,because we were able to amplify a 451-bp mRNAcorresponding to G3PDH from the worm (34). Sequence analysisof theMytilusPCR product demonstrated that them receptor fragmentexhibited 95% sequence identity with the human brainm opiate re-ceptor (32). Failure to demonstrate them receptor inAscaristissuessuggests that this material is intended for secretion into the microen-vironment as suggested by the MOR-IR localization.

DiscussionThe present study demonstrates for the first time the presence ofmorphine inA. suumand its expression under presumed nonstimu-lated (or basal) conditions. Animals maintained in the laboratoryfor 5 days also had morphine present and were secreting the ma-terial into the medium, demonstrating their ability to synthesizethis material. Unlike the free-living molluskMytilus edulis(32),Ascarisdoes not expressm opiate receptor transcripts. InAscaris,MOR-IR was found in the epidermis of the animal. These twopieces of data suggest that, inAscaris,morphine is secreted intothe microenvironment where it could be used as a signaling mol-ecule. We hypothesize that the morphine helps the parasite toevade the immunosurveillance machinery of the host and therebyincrease the stability of its microenvironment and ensure its sur-vival. This strategy appears to also ensure the survival of the ova.Furthermore, the morphineAscarisdisplays the same effect as nat-ural morphine on the NO release of the monocyte, an effect an-tagonized by the naloxone. This result suggests that the morphinefrom Ascariscan act on immune cells to enhance its survival.

Opioid peptides and their precursors are present in free-livingand parasitic invertebrates (21). These free-living invertebratescontain complex and highly specific opioid and opiate receptorsites (32, 35–37) involved in dopaminergic and NO processes (31,38). In these animals, the tissues that produced these signalingmolecules have the appropriate receptors to utilize them. For ex-ample, inMytilus, we recently demonstrated both morphine and am opiate receptor subtype that exhibited 95% sequence identitywith the human neuronalm opiate receptor (32). We had surmised,based on theMytilus model, that the signaling material was usedwithin that tissue. However,Ascarismakes morphine without acorresponding receptor, suggesting that it is secreted for “external”signaling, i.e., host immune down-regulation (8, 39). This hypoth-esis is supported by our immunohistochemistry results, whichdemonstrated the presence of this material in the epidermis of theanimal.

Complementing the present study are reports thatSchistosomasynthesize a morphine-like compound inhibiting immunocytes in analoxone-reversible manner (22). In this report, we also demon-strated that in worms maintained in the laboratory the endogenousmorphine-like levels were low and that, upon exposure to humanimmunocytes, a significant increase in the worm’s opiate leveloccurred, suggesting a feedback process. In the present report wealso note a significant drop in morphine levels in animals main-tained in the laboratory. However, its high level in the incubation

FIGURE 1. Purification of morphine fromAscaris. Morphine was iso-lated by HPLC. Running conditions: range, 5 nA; filter, 0.02 Hz; potential,500 mV; flow rate, 500ml/min; A buffer: 1 mM ammonium bicarbonate,10 mM ammonium chloride, 10 mM sodium chloride, 0.2 mM EDTA (pH5.0); B buffer: same as A buffer but with 50% acetonitrile. Gradient:t 5 0,0% buffer B;t 5 10 min, 10% buffer B;t 5 20 min, 30% buffer B;t 5 25min, 100% buffer B.A, Five nanograms of morphine internal standard.B,Standard curve of low morphine concentrations.C, Ascaris extract. D,Ascarisextract1 4 ng morphine external standard.E, Ascarisincubationmedium.

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medium indicates that as this material is made it is secreted into theimmediate environment. In this regard, if morphine were not syn-thesized in the worm, by day 5 it should not be detected becauseits tissue half-life is about 2 h and it cannot be detected in mam-malian and invertebrate tissues after 24 h (Ref. 40 and our unpub-lished observations).

TheseAscarismorphine data also suggest that morphine mayplay a role in gastrointestinal regulation. In human history it haswidely been accepted that diarrhea represents the body’s attempt toflush out toxins, including parasitic worms. The Persian physicianAvicenna prescribed morphine for cough, anemia, and diarrheathousands of years ago (41). Thus,Ascarismay secrete this opiate

FIGURE 2. Characterization ofAscarismorphine.A, GC/MS spectrum of two morphine standards (200 pg and 400 pg) and the morphine materialcollected during HPLC analysis ofAscaris.B, Bioassay of morphine-like activity extracted fromAscaris. Ascarisextracts stimulate NO release from humanmonocytes.Top,A total of 500ml of Ascariswhole body extract was added to a solution containing purified human monocytes (107 cells/ml) (arrowhead).This resulted in NO release.Bottom, Naloxone (first arrowhead; 1026 M) was added to another batch of cells under similar conditions before the additionof morphine extract (second arrowhead). This treatment blocked theAscarismorphine-stimulated NO release, demonstrating that theAscarismorphinematerial is opiate alkaloid in character.C andD, Immunocytochemical localization of morphine inAscaris. C,Ascarisbody wall region incubated withnonimmune 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 contractileparts of myoepithelial cells (d) (bar5 20 microns).E, Ascarisova incubated with nonimmune serum shows a complete lack of positive immunoreactivity.F, Morphine-like immunoreactivity immediately outside the ova (bar5 50 microns).

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alkaloid to diminish its “flushing” from the host because a sec-ondary effect on gastrointestinal motility is to induce constipation.

In conclusion, opioid processes appear to have evolved muchearlier than previously thought. Opiate alkaloid signaling appearsto have been conserved during evolution and used to enhance thelongevity of the host and parasite. Clearly, this represents an im-portant strategy for enhancing the odds for survival and, thus, forsimultaneously preserving the message.

AcknowledgmentsWe thank Danielle Benz, Ireen Khan, Mazen Madhoun (National Instituteof Mental Health-Career Opportunities Research Program Fellows), andYun Su Lee for technical assistance.

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FIGURE 3. Identification ofm opioid receptor mRNA transcript. TotalRNA isolated from these tissues was reverse transcribed, PCR amplified,and run on agarose gels.Lane 1, G3PDH housekeeping gene (expectedtranscript size is 451 bp).Lane 2,m opioid receptor transcript inMytiluspedal ganglia (expected size, 441 bp).Lane 3, lack ofm receptor transcriptin Ascaris. Lane 4, 100-bp DNA markers. Both negative and positivestrands were sequenced; however, only the sequence from the negativestrand is shown inlane 2. The tissues were washed extensively with PBSto limit any contamination.

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