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Journal of Insect Physiology 52 (2006) 870–880

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Structure and function of the extraembryonic membrane persistingaround the larvae of the parasitoid Toxoneuron nigriceps

A. Grimaldia,�, S. Cacciab, T. Congiuc, R. Ferraresea, G. Tettamantia, M. Rivas-Penaa,G. Perlettia, R. Valvassoria, B. Giordanab, P. Falabellad, F. Pennacchiod, M. de Eguileora

aDipartimento di Biologia Strutturale e Funzionale, Universita dell’Insubria, via Dunant 3, 21100 Varese, ItalybDipartimento di Biologia, Universita di Milano, via Celoria 26, 20125 Milano, Italy

cDipartimento di Biologia, Difesa e Biotecnologie Agro-Forestali, Universita della Basilicata, Viale dell’Ateneo Lucano 10, 85100 Potenza, ItalydDipartimento di Entomologia e Zoologia Agraria ‘‘F. Silvestri’’, Universita di Napoli ‘‘Federico II’’, Via Universita 100, 80055 Portici (NA), Italy

Received 1 February 2006; received in revised form 16 May 2006; accepted 16 May 2006

Abstract

The embryo of Toxoneuron nigriceps (Hymenoptera, Braconidae) is surrounded by an extraembryonic membrane, which, at hatching,

releases teratocytes and gives rise to a cell layer embedding the body of the 1st instar larva. This cell layer was studied at different

developmental times, from soon after hatching up to the first larval moult, in order to elucidate its ultrastructural, immunocytochemical

and physiological function. The persisting ‘‘larval serosa’’ shows a striking structural and functional complexity: it is a multifunctional

barrier with protective properties, limits the passage of macromolecules and it is actively involved in the enzymatic processing and uptake

of nutrients. The reported results emphasizes the important role that the embryo-derived host regulation factors may have in parasitism

success in Hymenoptera koinobionts.

r 2006 Elsevier Ltd. All rights reserved.

Keywords: Serosal membrane; Host regulation; Larval development; Nutrition; Sugar absorption; Immunoevasion

1. Introduction

Embryonic development in parasitic Hymenoptera hasbeen studied in relatively few species but from what weknow, it appears that the extraembryonic membranes havedifferent developmental origins in different species (Trem-blay and Caltagirone, 1973). The formation duringembryogenesis may involve the polar bodies, thoughdifferent origins have been proposed for denoting theseextraembryonic membranes (Tremblay and Caltagirone,1973; Schmidt, 2000; Lamer and Dorn, 2001). The term‘‘serosa’’ has been largely used in most cases, even to definenon-homologous extraembryonic envelopes.

Whatever the origin of these extraembryonic membranesin parasitic Hymenoptera, their structural organizationstrongly suggests that they have nutritional and protectiveroles (Ivanova-Kasas, 1972; Tremblay and Caltagirone,

e front matter r 2006 Elsevier Ltd. All rights reserved.

sphys.2006.05.011

ing author. Tel.: +390332 421312; fax: +39 0332 421300.

ess: annalisa.grimaldi@uninsubria.it (A. Grimaldi).

1973; Dorn, 1976; Dyby and Silhacek, 1997; Fausto et al.,2001; Lamer and Dorn, 2001). For example, Copidosoma

floridanum, has a passive immunoevasion strategymediated by a polar body-derived extraembryonic mem-brane (Corley and Strand, 2003).When the parasitoid larva hatches, the extraembryonic

membranes may dissociate in isolated cells freely floatinginto the host haemocoel, currently defined as teratocytes(see for review Dahlman and Vinson, 1993), or they mayremain undissociated or partially dissociated (Grandori,1911; Hawlitzky, 1972; Strand et al., 1986; Lawrence, 1990;Volkoff and Colazza, 1992; Pennacchio et al., 1994;Beckage and de Buron, 1994, Corley and Strand, 2003;Pedata et al., 2003).The role of teratocytes in the host regulation process has

been studied in a limited number of braconid species. Thesefunctional studies have described different effects on hostnutritional suitability, endocrine balance and immuneresponse (see for review Dahlman and Vinson, 1993;Beckage and Gelman, 2004; Pennacchio and Strand, 2006).

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In contrast limited information is available on thephysiological functions of undissociated or partiallydissociated extraembryonic membranes (Lawrence, 1990;Corley and Strand, 2003).

Toxoneuron nigriceps (Hymenoptera, Braconidae) is anendophagous braconid attacking the larvae of the tobaccobudworm, Heliothis virescens (Lepidoptera, Noctuidae).The embryo of this parasitoid species is surrounded by anextraembryonic membrane, denoted as serosa, from whichteratocytes originate by dissociation of the anterior andposterior poles (Pennacchio et al., 1994). The releasedteratocytes, along with the associated polydnavirus, areresponsible for the endocrine disruption of the host(Pennacchio et al., 1997, 2001), whilst the effective role ofthe persisting serosal layer around the newly hatched larvaestill remaines elusive.

To fill this gap, the serosa, which surrounds the 1st instarlarvae of T. nigriceps as a continuous cell layer, was studiedat different developmental times, from soon after hatchingup to the first larval moult by performing an ultrastructur-al, immunocytochemical and physiological characteriza-tion. These observations indicate that the persisting ‘‘larvalserosa’’ may play an important role in evasion of the hostimmune response, act as a barrier for macromolecules, andformation in hydrolyse and absorption of nutrients.

2. Materials and methods

2.1. Insect rearing

The braconid parasitoid T. nigriceps was reared onH. virescens larvae according to Vinson and coworkers(1974). Rearing temperature for both parasitoid adults andparasitized host larvae was 2970.5 1C, and the photoper-iodic regime was L16:D8.

2.2. Host parasitization and dissection

Host larvae were singly parasitized as late 4th instars byexposing them to a parasitoid female in a rearing vial. Onlyone oviposition attack was allowed, and larvae that wereaccidentally superparasitized were discarded.

At fixed time intervals after parasitization, H. virescens

larvae were CO2 anaesthetized and dissected in Pringle’ssaline (1938), to explant 1st instar parasitoid larvae, atdifferent time points of their development. The recoveredparasitoid larva was immediately transferred in a drop ofsterile saline onto a microscope slide and covered with acoverslip. The specimen was then observed and photo-graphed using an Olympus BH2 light microscope,equipped with a Leica DC 300F camera.

2.3. Light microscopy, scanning electron microscopy

(SEM) and transmission electron microscopy (TEM)

T. nigriceps 1st instar larvae were extracted fromparasitized host larvae as described above and fixed in

2% glutaraldehyde (in 0.1M Na-cacodylate buffer, pH7.2), for 2 h at room temperature. Specimens were thenpost-fixed for 2 h with 1% osmic acid in 0.1M Na-cacodylate buffer at room temperature, and dehydrated inan ethanol series.For transmission electron microscopy (TEM) prepara-

tion, dehydrated samples were embedded in an Epon-Araldite 812 mixture and sections were cut with a ReichertUltracut S ultratome (Leica, Nussolch, Germany). Semi-thin sections were stained by conventional methods (crystalviolet and basic fuchsin) and observed with a lightmicroscope (Olympus, Tokyo, Japan). Images wereacquired with a Leica DC 300F camera.Thin sections were stained with uranyl acetate and lead

citrate, and observed with a Jeol 1010 EX electronmicroscope (Jeol, Tokyo, Japan).For scanning electron microscopy (SEM) preparation,

dehydrated samples were dried according to Congiu et al.(2004), mounted on stubs, gold coated with a Sputter K250coater (Emitech), and observed with a SEM-FEG XL-30microscope (Philips).

2.4. Alcian Blue staining

Selective staining of glyco-conjugated compounds wascarried out according to Tettamanti et al. (2004). Tissuesamples of T. nigriceps larvae were fixed for 3 h at 4 1C,with 3% glutaraldehyde in phosphate buffered saline (PBS)0.1M, pH 7.4, and then cut into small blocks of about100 mm. After a 1 h preincubation at room temperature, in25mM MgCl2–acetate buffer (pH 5.8), samples wereincubated for 7 h in the staining solution (0.05% w/vAlcian Blue 8 GX in 25mM acetate buffer and MgCl2 atfinal concentrations of 0.3M). Then, samples were washedfor 1 h in MgCl2-acetate buffer, for 4 h in 0.01N HCl, for1 h in distilled water and finally, in PBS for 30min.Dehydration in an ethanol series preceded inclusion inEpon resin. Control sections were fixed and processed asabove but in the absence of Alcian blue.

2.5. Immunohistochemistry

First instar larvae of T. nigriceps explanted fromanesthetized H. virescens larvae were dissected and fixed,according to Causton (1984), in 4% paraformaldehydesolution in 0.2M PBS (pH 7.2) containing 0.5% glutar-aldehyde for 2 h at 4 1C. The specimens were then washedin the same buffer and, after a standard step of serialethanol dehydration, were embedded in Epon-Araldite, asdescribed above, and sectioned with a Reichert Ultracut Sultratome. After etching with 3% NaOH in absoluteethanol, 0.7 mm sections, were incubated for 30min withPBS containing 2% bovine serum albumin (BSA). Semi-thin sections were treated for 2min with a resin-removingmixture (2 g KOH in 5ml propylene oxide and 10mlmethanol), rinsed with methanol and placed in PBS.Sections were incubated for 30min with the following

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primary antibodies (dilution 1:200): monoclonal anti collagenIV (Novocastra Laboratories, Newcastle upon Tyne, UK)and polyclonal anti-GLUT2 (Glucose Transporter 2) (Che-micon International Inc., CA, USA). Specimens were washedand incubated with the appropriate fluorescein isothiocyanate(FITC)-conjugated secondary antibodies. Afterwards, sec-tions were rinsed in distilled water and counterstained withcrystal violet. The PBS buffer used for the washing steps andantibody dilutions contained 2% BSA.

Control experiments were performed by omitting theincubation with the primary antibody. Semithin sectionswere collected on slides and coverslips mounted inVectashield mounting medium for fluorescence. Slideswere then examined with a BIORAD confocal microscope.

2.6. Histochemical detection and enzymatic activity of

trehalase

For histochemistry first instar larvae of T. nigriceps

explanted from anesthetized H. virescens larvae wereembedded in Poly-freeze tissue freezing medium (Poly-sciences, Eppelheim, Germany) and immediately frozen inliquid nitrogen according to Geiger et al. (1980). Cryosec-tions (7 mm) were collected on slides and treated accordingto Pearse (1989) for trehalase visualization. Mounted slideswere observed by light microscopy.

The parasitoid larva lives in the H. virescens haemolymphrich in trehalose. Thus the ability of T. nigriceps larvae tohydrolyze trehalose was assessed by measuring the amount ofglucose released after 1h of incubation at room temperature(22–24 1C), in 200ml of the physiological solution used forglucose uptake measurements (see next section). Incubationwas carried out with 30 larvae, when 1st instars were used,while in the case of 2nd and 3rd instars a single larva wassufficient to detect the released glucose by means of anenzymatic colorimetric test (Glucose Liquid, Sentinel, CH).Trehalase (E.C.3.2.1.28) activity was expressed in mU/larvaand mU/mg of larval proteins, considering that 1 unit (U) ofenzyme hydrolyzes 1mmol of substrate/min. To measure theamount of larval proteins, at the end of the incubation,parasitoid larvae were dissolved in 50ml of NaOH 1N at roomtemperature for 1h, for early and late 1st instar, or 4h for the2nd and 3rd instars. Protein concentration was measured intriplicate in 1ml sample aliquots, according to Bradford (1976),using the Coomassie Brilliant Blue G-250 protein assay (Pierce,Rockford, IL, USA) and bovine serum albumin as standard.

After removal of the larvae, the incubation medium wasleft at room temperature for 1 h. Then, trehalase activitywas measured again to assess if any enzyme had beenreleased in the medium. The mean values of the enzymaticactivity measured at the beginning and after 1 h werecompared by Student’s t test.

2.7. Glucose uptake in vitro

Incubations of synchronized T. nigriceps early 1st instarlarvae were performed at room temperature (22–24 1C) in

the following physiological solution (in mM): 5 K2SO4, 1NaCl, 5 MgSO4, 1 CaCl2, 10 K-citrate, 240 trehalose, 5Tris–HCl at pH 6.8. This solution mimics the basic ioniccomposition, osmolarity and pH value of lepidopteranlarvae haemolymph. Glucose absorption across larvalserosal cells and epidermal epithelium was determined bymeasuring the uptake of radiolabeled 3-O-Methyl-D-Glucose (3-MG), a glucose analogue that is not metabo-lised but recognized by GLUT2-like glucose transporters inAphidius ervi larvae (Giordana et al., 2003). Three to fourlarvae were incubated for 15min in 300 ml of physiologicalsolution containing 0.1mM 3-O-Methyl-D-[1-3H]Glucose(30 mCi/ml) (Amersham Biosciences, Italy). The experi-mental larvae were preincubated for 30min in thephysiological solution above in presence of the inhibitorcytochalasin B (0.2mM) (Sigma-Aldrich, Italy), prior tothe addition of labeled 3-MG. At the end of incubationsthe radiolabeled medium was removed, the larvae wererinsed 3 times with 2ml of physiological solution and thentransferred to an eppendorf vial containing 100 ml of 80%ethanol (v:v) in water. The vial was left for 24 h at roomtemperature, then centrifuged for 3min at 10,000 g, thesupernatant was recovered and processed for liquidscintillation counting (Packard Tri-Carb 1600CA). Theethanol-insoluble pellet was dissolved in 50 ml of NaOH1N for 60min at room temperature. Protein concentrationwas measured in triplicate in 1 ml sample aliquots accordingto Bradford (1976), with the Coomassie Brilliant Blue G-250 protein assay (Pierce, Rockford, IL, USA) and BSA asstandard.The mean values of the experimental uptakes were

compared by Student’s t test.

2.8. Localization of transported 3-MG by autoradiography

Early 1st instar larvae were incubated for 15min in thephysiological solution containing 0.1mM 3-O-Methyl-D-[1-3H] Glucose. The radiollabeled solution was removed,the larvae were rinsed 3 times, then recovered andimmediately frozen in liquid nitrogen. Frozen sections,7 mm thick, were collected on slides. Slides were coated withKodak fine-grain Autoradiography Stripping Plate AR 10emulsion, air-dried overnight in the dark, and then storedat 4 1C for 2 weeks in an airtight–lightproof box with silicagel desiccator. A Kodak D-19 developer was used. Theslides were stained with hematoxylin and mounted inPermount. Controls were exposed to the radiolabelledsolution for 1min and then processed as described above.

2.9. Analysis of the permeability to macromolecules of the

basal membrane

Parasitized H. virescens larvae were anesthetized andeach larva was injected with dextran solution in PBS(1 mg/ml). Three types of dextrans were tested: neutral3 kDa dextrans Texas Red-conjugated, anionic 3 kDadextrans FITC-conjugated (Molecular Probes, OR, USA)

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and neutral 250 kDa dextrans FITC-conjugated (SIGMA,MO, USA). Twenty microliters of each type of dextransolution were injected, as a single dose, into the haemocoel.One hour after injection, H. virescens larvae wereanesthetized and dissected to explant 1st instar parasitoidlarvae, which were embedded in Poly-freeze tissue freezingmedium (Polysciences, Eppelheim, Germany) and immedi-ately frozen in liquid nitrogen. Controls were not exposedto fluorescent molecules.

Cryosections (7 mm) were collected on slides and cover-slips mounted in Vectashield mounting medium forfluorescence. Slides were then examined with a BIORADconfocal microscope.

Another group of parasitized H. virescens larvae wasinjected and fixed in 4.1% glutaraldehyde in 0.1Mcacodylate buffer (pH 7,4) with the addition of 2.5%tannic acid (1.7 kDa in size). One hour after injection theexplanted parasitoid larvae were prepared for transmissionelectron microscopy as previously described.

Tannic acid has been used for delineating the extra-cellular compartment and detecting permeability of cellsand/or extracellular matrix.

3. Results

3.1. Morphological and molecular characterization of the

serosa surrounding T. nigriceps larvae

Early 1st instar larvae of T. nigriceps were 1.5–2mmlong, showed sclerotized head capsule and mandibles, awell segmented body which contained a large gut, and wasentirely surrounded by a thick serosa, as can be observed inlarval whole mounts (Fig. 1A), and in samples processedfor optical (Figs. 1B, C) or SEM (Fig. 1D) observations.This serosal envelop was mainly composed of partiallyoverlapping large cells (Fig. 1C). At higher magnification,serosal cells (Fig. 1E) showed a cytoplasm with many freeribosomes, numerous small mitochondria, an abundance ofrough and smooth endoplasmic reticulum, and large andirregular nuclei (Fig. 1F). Each serosal cell displayed aplasma membrane showing the ubiquitous presence ofmicrovilli, organised in a brush border (Figs. 1E,G–J, 2A).

Serosal cells showed plasma membrane domainswith morphological differences. The surface bathed byH. virescens haemolymph was characterized by a brushborder covered by a thick amorphous felt (Figs. 1E,G–I).On the lateral sides of the cell, the microvilli were regularlyspaced and intermingled with those of the adjacentcells (Figs. 1E,G, J). The area facing the surface of theparasitoid body presented crowded microvilli (Figs. 2A, B).A thick and uninterrupted fibrillar basal membrane waspresent between the parasitoid body and the serosal cells(Figs. 2B–F). The coupling among the basal membrane andthe plasma membranes of two adjacent serosal cells formedcharacteristic deep and convoluted ‘‘sandwich-like’’ struc-tures shown in Figs. 2C–F.

Collagen IV was abundantly present in the thick basalmembrane (Figs. 2G–I).Alcian Blue is a specific stain used for the visualization of

glyco-conjugated substances, such as proteoglycans. Theexternal felt, the surfaces of the microvilli on the serosalcell brush border, and the basal membrane coating thelarval body wall, showed an intense and uniform reactionsignal (Figs. 1I, 2E) which was not evident in controlsections (Figs. 1H, 2C).In late 1st instar larvae, the stretched ‘‘larval serosa’’ was

thinner than in early 1st instar larvae and no longer incontact with both the surface of the larval body and theexternal amorphous felt. The cells showed many largevacuoles and the presence in the cytoplasm of severalbacteria or residues of bacterial cells (Figs. 3A, B). Fullygrown 1st instar larvae, ready to moult, showed serosalenvelope characterized by a reduced thickness (Fig. 3C), alarger space between the external microvilli and the felt(Figs. 3C–E), while adjacent cells were no longer tightlyconnected as the lateral microvilli disentangled (Fig. 3F).

3.2. Nutritional function of the serosa surrounding

T. nigriceps larvae

A possible role of the serosa as a nutritional interfacebetween the embryo or 1st instar larvae of T. nigriceps andthe surrounding host environment was suggested byPennacchio et al. (1994). The demonstration of nutrientabsorption, such as glucose and amino acids, through thebody surface by the larvae of another braconid parasitoid,A. ervi (Giordana et al., 2003; Caccia et al., 2005),stimulated the idea of assessing if a similar phenomenonoccurred also in T. nigriceps larvae and understanding therole of the ‘‘larval serosa’’, if any, in this process.

T. nigriceps larvae are bathed by the host haemolymphrich in trehalose which, to be used as a source of glucose,has to be cleaved by a trehalase (a,a—trehalose gluco-hydrolase). Then, we investigated if the external surface ofparasitoid larvae (at all instars) had any trehalase activityand, if so, its localization. We sought to better understandthe possible modulation/changes of this enzymatic activityduring larval development.The enzyme was located in the plasma membranes both

of serosal and epidermal cells (Figs. 3G–K). The amount ofglucose released by T. nigriceps larvae incubated in atrehalose-rich solution varied considerably during larvaldevelopment. The total trehalase activity per larvaincreased with body mass, from 1st to 3rd instar(Figs. 3L,M) but the specific enzymatic activity, expressedper mg of larval proteins, was comparatively high alreadyin 1st instar larvae, with a peak corresponding to the serosadisruption occurring in fully grown larvae ready to moult(Fig. 3M). At that stage significant trehalase activity wasdetected in the incubation medium 1h after the larvae hadbeen removed, likely due to serosal residues being shed intothe medium. The enzyme was not released by early 1st, 2ndand 3rd instar larvae, because the amount of glucose did

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Fig. 1. (A–C) Light microscopy. Early 1st instar larvae of T. nigriceps: (A), live mounted sample showing the characteristic mandibles (m), and semithin

sections of the body of the larva (p) surrounded by a thick serosa (encircled area, s) (A–C), made of partially overlapped cells (C). Scale bars: (A) 300mm;

(B) 20mm; (C) 50 mm. (D–J): SEM (D), TEM (E, F, H, I, J) photographs and schematic drawing (G) of the larval serosa. The serosal cells are characterized

by irregular nuclei (F) and ubiquitous brush border (E, G–J). Microvilli are visible on the external surface, where are covered by an amorphous felt (f)

(E, G–I) and on the lateral cell side (E, G, J). The external amorphous felt and the microvilli are Alcian Blue positive (I). The positivity of the reaction

(dark color) is a specific visualization of glyco-conjugated substances such as proteoglycans. Scale bars: (D) 70 mm; (E) 3.5 mm; (F) 6mm; (H) 1.4mm;

(I) 1.0 mm; (J) 1.0mm.

A. Grimaldi et al. / Journal of Insect Physiology 52 (2006) 870–880874

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Fig. 2. (A–F) TEM photographs (A, C, D, E, F) of the encircled area in the schematic drawing (B). Regularly spaced microvilli close to the surface of the

parasitoid (p) are visible (A). The thick fibrillar basal membrane (bm) lines the parasitoid surface (C) and forms characteristic ‘‘sandwich-like’’ structures

(D–F) (arrowheads). The coupling among the basal membrane and the plasma membranes of two adjacent serosal cells is clearly evident (D–F). Scale bars:

(A) 0.8 mm; (C) 0.3mm; (D) 1.8 mm; (E) 1.0mm; (F) 0.2 mm. (G–H) light microscopy. T. nigriceps serosa (s) (G). Immunostaining of serosa (s) using anti-

collagen IV mAb: the basal membrane is strongly positive (H). Control section (I).

A. Grimaldi et al. / Journal of Insect Physiology 52 (2006) 870–880 875

not increase in the incubation solution after removing theexperimental larvae.

The ability of the parasitoid larva to absorb the releasedglucose was assessed by measuring the uptake in early 1st

instar larvae of the glucose analogue 3-MG. This analogue,which is not metabolized, was absorbed and the transportwas reduced by 50% in the presence of cytochalasin B(Table 1), a specific inhibitor of the facilitative glucose

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Fig. 3. (A–F) TEM (A, B, D, E, F) and light microscopy photographs (C) of late 1st instar larvae of T. nigriceps (p). The cytoplasm of serosal cell is

occupied by large vacuoles (A, D, E) and by bacteria (A, B) (arrowheads). The stretching and detachement of serosal cells (s) is evident both in

correspondence of the parasitoid body surface (C–E) and between adjacent cells (F). Scale bars: (A) 5 mm; (B) 1.6mm, (C) 70 mm, (D) 3.5mm; (E) 10 mm; (F)

3.8mm. (G–K):Cryosections of 1st (G–I), 2nd (J) and 3rd (K) instar larvae. An intense enzymatic activity is detected in all larval stages. The signal is

localized both in the serosal (encircled areas, s) and epithelial cells (e) of 1st instar larvae (H, I). Control section (G). In 2nd and 3rd instar larvae the serosa

is lost and the enzyme is localized in epithelial cells (e) of the parasitoid (p) (J, K). (L–M): Threalase activity. In 1st instar larvae, the serosal cell layer

contributes to the digestive activity, a contribution that becomes particularly important in the last part of the instar, when the breakdown of the tissue and

the cell disruption cause a release of the enzyme in the external microenvironment, enhancing considerably the total enzyme activity.

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transporters GLUT1-4 (Uldry and Thorens, 2004). Inlarvae incubated for 15min with radiolabeled 3-MG andanalyzed by autoradiography, the silver particles wereclearly visible in the cytoplasm of serosal cells, in theepidermal cell layer and in the larval tissues underneath theepidermis (Fig. 4A). The commercial antibody raisedagainst the mammalian glucose transporter GLUT2 gaverise, in early 1st instar larvae, to a strong cross-reactionsignal uniformly distributed along the plasma membrane ofthe serosal cells (Figs. 4B–D).

3.3. The permeability of the basal membrane

In order to characterize permeability of the serosa,molecules with different electric charge and molecular masswere used. Dextrans with a molecular mass of 3 kDa,independently of their positive charge, crossed the serosalenvelope (Figs. 4E–G) while 250 kDa dextrans wereblocked (Fig. 4H).

The small tannic acid molecules (1.7 kDa), negativelycharged, were observed among adjacent serosal cells (Figs.4I–L), but they were blocked and accumulated along theconvoluted basal membrane (Fig. 4M).

4. Discussion

The cell layer of serosal origin ‘‘glued’’ to the body of 1stinstar larvae of T. nigriceps is a multifunctional barrier,that limits the passage of macromolecules, is activelyinvolved in the enzymatic processing and uptake ofnutrients and likely performs a protective role against hostimmune reaction.

The inner side of this serosal layer, facing the larvalbody, shows a complex organization with peculiar mor-phological features that are surprisingly similar to thosepresent in vertebrate kidneys, where the filtration barrier isformed by the double-sided glomerular basal laminaoccurring between the endothelial cells and the podocytes(Osawa et al., 1966). The basal membrane, coating thelarval body, primarily formed by collagen IV andproteoglycans (the main components of both invertebratesand vertebrates basal membranes), shows deep invagina-tions between plasma membranes of adjacent serosal cells,enhancing cell-cell interaction, as also evidenced in othertissues (Pease, 1956). These sandwich-like structures,mediating stronger adhesion between cells and the basalmembrane, form a continuous inner belt which operates as

Table 1

Uptake of 3-O-Methyl-D-Glucose by early 1st instar T. nigriceps larvae

and inhibition by 0.2mM Cytochalasin B

Uptake (pmol/mg protein/30min)

Control 326755 (7)

Cytochalasin B 148723 (4)�

Mean7SE, number of experiments in parenthesis.�Po0.05.

a filtration barrier, blocking the free diffusion of largemacromolecules such as fluoresceinated dextrans (about250 kDa) or low molecular weight molecules bearingnegative charges like tannic acid (1.7 kDa), while it allowsthe passage of low molecular weight molecules likefluoresceinated dextrans of about 3 kDa, independently ofthe presence of a positive charge. These results indicate thatthe negative fixed charges, conferred to the basal mem-brane by proteoglycans and collagen IV, influence thepassage of circulating molecular components present in thehost haemolymph that bathes the parasitoid serosa.In addition the proteoglycans that are present on the

surface of microvilli and in the external felt surroundingthe serosa, are among the main components of basalmembranes of insects. The basal membrane, in insects,lines the haemocel and all the internal organs, and thecirculating haemocytes normally do not attach or attachweakly to this substrate, as long as it remains unaltered(reviewed in Lavine and Strand, 2002). Then, it is likelythat the presence around the parasitoid larval body ofmultiple layers containing molecular components abun-dantly present in the basal membrane may provideprotection against encapsulation, especially if these com-ponents are similar to those found in the basal membranesof the host.The presence of bacterial cells or cell residues in the

serosal layer around the parasitoid larval body suggeststhat these cells of embryonic origin provide some protec-tion also against potential pathogens, when the immunesystem of parasitized hosts is compromised (Ferrarese etal., 2005); the early expression of TnBV encoded proteinsimpairs the immune response against both parasites andpathogens (Falabella et al., 2006). The concurrent presenceof active and passive immunoevasive strategies, even at theegg–embryo stage, which has an external fibrous layerconferring protection against the host immune response(Vinson and Scott, 1974), indicates that in this host–par-asitoid association, like in many others, several mechan-isms are concurrently active to better insure survival of theprogeny.The ultrastructural observations clearly support our

hypothesis that the cells of ‘‘larval serosa’’ are activelyinvolved in intense nutritional exchanges as stronglysuggested by the thick array of microvilli, which arenormally associated with absorbing functions. Further-more, like in a gut epithelium, this structural organization,resulting in a considerable extension of the active surface, isassociated with intense enzymatic activity. The parasitoidlarva absorbs glucose, obtained from enzymatic processingof trehalose, which is the most highly abundant sugar inthe host haemolymph. The histochemical characterizationof the parasitoid, assessing the presence of trehalaseactivity associated with the serosa, and the trehalaseactivity detected in the incubation medium of late 1stinstar larvae loosing their serosal cells, collectively demon-strate that this cell layer of embryonic origin is able tohydrolyze trehalose. In later larval stages (2nd and 3rd), the

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Fig. 4. (A) Autoradiography (A) of early 1st instar larvae surrounded by serosa (square bracket), with 3-0-methyl-D-[1-14H]- glucose. Labelled granules

are visible in the serosal cells (s) and in the larval tissues of parasitoid (p), indicating the occurrence of intense glucose uptake. (B–D) Semithin sections of

early 1st instar larvae (B). Immunodetection of Glucose Transporter 2 (GLUT2) localized along the plasma membrane of each serosal cells (C).

D Negative control section. (E–H) Cryosections of T. nigriceps early 1st instar larvae (E–H), explanted from host larvae into which three types of dextrans

were injected: anionic 3 kDa dextrans (FITC-conjugated) (E) and neutral dextrans with different molecular weight such as 3 kDa, Texas red-conjugated

(F), and 250 kDa, FITC-conjugated (H). The 3 kDa dextrans cross the serosal membrane, independently of their charge (E, F), while large molecules (H)

are blocked and accumulated in the middle part of serosa (s, square brackets). (G) Negative control. (I–M) TEM photographs of T. nigriceps early 1st

instar larvae (I–M), explanted from host larvae into which tannic acid, 1.7 kDa in size and negatively charged, was injected. The positivity of the reaction

(dark spots) is localized in the extracellular matrix among cells (I), homogeneously distributed if the section plane is close to the surface facing the

haemolymph (i.e. it does not involve the invagination areas close to the parasitoid body). Dark spots are visible on and within the superficial amorphous

felt (f) (J) and close to the lateral microvilli (K, L) while signal is not present in the convolute basal membrane (bm) (M). Scale bars; (I) 8mm, (J) 5.0mm;

(K) 5.0mm; (L) 2 mm; and (M) 2 mm.

A. Grimaldi et al. / Journal of Insect Physiology 52 (2006) 870–880878

ARTICLE IN PRESSA. Grimaldi et al. / Journal of Insect Physiology 52 (2006) 870–880 879

trehalase activity is present in the epidermal cells and, likein the serosal cells, is associated with the facilitative glucosetransporters of the GLUT family (data not shown). Wemay hypothesize that in the early 1st instar larvae, theserosal cells, with their high number of interdigitatedmicrovilli, can create a microenvironment where theconcentration of glucose released by trehalase activity ismaintained at high levels. This glucose concentrationgradient can provide the driving force for its uptake bythe serosal and epidermal cells, mediated by the facilitativeglucose transporters. Indeed, the glucose analogue 3MGwas readily absorbed by T. nigriceps larvae (Table 1), andits uptake was inhibited by cytochalasin B, which indicatedthe occurrence of GLUT2 transporters (Caccia et al.,2005). The autoradiography of incubated larvae clearlyshowed that a large uptake of glucose was also performedby the serosal cells to sustain their metabolism. Thisabsorbing capacity of the serosal cells around the larvalbody may also be the residual activity of a more intenseabsorption pathway of the embryonic serosa, in support ofthe growing embryo. The overall importance of thismechanism for the nutrition of T. nigriceps immaturestages is further supported by the precocious increase oftrehalose titre registered in parasitized H. virescens larvae(Vinson, 1990). The presence and persistance of a serosallayer around the body of 1st instar larvae, facilitating theuse of host sugars, indicates that the early stages ofdevelopment may need a more substantial carbohydrateintake to support their growth and development. This hasto be taken into account to further develop the artificialdiets defined for this endophagous parasitoid (Pennacchioet al., 1992).

The finding that glucose is largely absorbed by theepidermal cells and then transferred within the larval bodyis not new, as we demonstrated that it occurs also in thelarvae of another braconid species A. ervi, parasitoid ofaphids (de Eguileor et al., 2001; Giordana et al., 2003).Even though this phenomenon has been only recentlydemonstrated, based on morphological and functionalobservations, Grandori (1911) postulated about a centuryago its occurrence in Cotesia glomerata, a species that alsoshows the persistence of the extraembryonic membranearound the larva (Grandori, 1911: Beckage and de Buron,1994).

In conclusion, the experimental data we report in thispaper indicate that the serosal layer persisting around thebody of 1st instar larvae of T. nigriceps is a complexstructure that likely contributes both to the immuneevasion and to the nutritional exploitation of the host.This further emphasizes the important role that theembryo-derived host regulation factors may have inparasitism success in Hymenoptera koinobionts.

Acknowledgements

We are grateful to Maria Luisa Guidali for technicalassistance and to Kathleen Histon for revising the English.

In addition, the authors would like to thank ProfessorOttaviani and Professor Raspanti for their critical discus-sion.This work has been supported by MIUR-FIRB-COFIN

Grants no. RBNE01YXA8/2004077251 and by CentroGrandi Attrezzature of University of Insubria.

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