InternarionalJournnlforParasirology Vol. 17, No. 7,pp. 1257-1265, 19X1

Pnnfed in Grear Britain.

0020-7519/X7 $3.00 + 0.00 Pergamon Journals Lrd.

0 1987 Ausrrahan Socray for I’arasiro~ogy.

LIGHT AND ELECTRON MICROSCOPICAL STUDIES OF THE LOCATION OF STRONGYLOIDES ST’RCORALIS IN THE JEJUNUM OF

THE IMMUNOSUPPRESSED DOG

D. I. GROVE,* A. WARTON ,t L. L. Yu,t C. NORTHERN* and J. M. PAPADIMITRIOU~

Departments of *Medicine and tPathology, University of Western Australia, Queen Elizabeth II Medical Centre, Nedlands, Western Australia, 6009

(Received 11 June 1986)

Abstract-GRovE D. I., WARTON A., Yu L. L., NORTHERN C. and PAPADIMITRIOU J. M. 1987. Light and electron microscopical studies of the location of Strongyloides stercoralis in the jejunum of the immuno- suppressed dog. International Journal for Parasitology 17: 1257-1265. This study was undertaken to define the precise anatomical location of .S. stercoralis in the intestinal mucosa of the dog. In order to facilitate finding worms, hyperinfection was induced by immunosuppression of the host with prednisolone. Light microscopy showed adult worms in vacuoles in close relationship with the columnar epithelium although portions of worms were sometimes seen in the intestinal lumen. Electron microscopy demonstrated that adult worms were situated between the enterocytes. The enterocytes were compressed and distorted and appeared to form a tunnel through which the worm probably moved. Adult worms were never observed to penetrate the basement lamina and enter the lamina propria. Rhabditiform and filariform larvae were never seen in the mucosa. These data suggest that autoinfection does not occur by rhabditiform larvae being seeded directly into the tissues, but supports the hypothesis that they are deposited in the small bowel lumen, moult, then penetrate the mucosa of the lower gastrointestinal tract. Since adult worms dwell in the epithelial layer of the mucosa, they are susceptible to attack by cellular elements of the host’s defences, although this was not observed in the present study because the host was immunosuppressed.

INDEX KEY WORDS: Strongyloides stercorulis; dog; immunosuppression; electron microscopy.

INTRODUCTION

NINE YEARS after the discovery of Strongyloides stercoralis by Normand in 1876, Golgi and Monti reported that the adult worms were located in the crypts of Lieberkuhn in the small intestine (Golgi & Monti, 1885). Several years later, Sonsino reported finding eggs and larvae, not only in the depths of the crypts, but also in the mucosa (Sonsino, 1891). In 1900, Askanazy indicated that he had found Strongy- Zoides infection during the course of a post-mortem examination of a patient who had died from metastatic cancer of the lung. He asked the rhetorical question:

“What relationship does A. intestinalis [Anguillula intes- tinulis = S stercorulis] have to the intestinal wall?” (Askanazy, 1900)

Remarking that the muscularis mucosae usually

checked the further penetration of the parasite, he

answered his question thus:

Correspondence: Dr. D. I. Grove, Department of Medi- cine, University of Western Australia, Queen Elizabeth II Medical Centre, Nedlands, Western Australia 6009.

“The A. infestinulis bores into the intestinal wall, primarily into the mucosa, and often into the epithelium.” (Askanazy, 1900)

Confusion as to the precise location of the parasite in the small intestine has led to a succession of reviewers merely stating that S. stercorulis invades the intestinal mucosa (Belding, 1942; Faust, Russell & Jung, 1970; Manson-Bahr & Apted, 1982; Gilles, 1984; Mae- graith, 1984; Markell, 1985).

Similar uncertainty has surrounded the location of S. ratti in the intestinal mucosa of rodents. Some authors have considered that worms invaded and developed in the gut mucosa (Abadie, 1963; Wert- heim & Lengy, 1965), whereas others were of the view that adult worms were located deep in the lumen of the crypts and did not penetrate the epithelium (Genta & Ward, 1980). We have shown recently by electron microscopical studies of mice that S. rutti adult worms are situated between the enterocytes and that neither they nor rhabditiform larvae penetrate deep into the basement lamina (Dawkins, Robertson, Papadimitriou & Grove, 1983).

Strongyloides stercoralis differs from S. ratti in that it has the capacity to autoinfect. The mechanisms of

1257

12.58 D. I. GROVE, A. WARTON, L. L. Yu, C. NORTHERN and J. M. PAPADIMITRIOU

autoinfection are obscure and it is conceivable that the structural nature of the host-parasite interaction may play a part in the genesis of this phenomenon. We have therefore used electron microscopical techniques to examine the precise location of S. stercoralis in the small intestine of the dog. In order to facilitate the finding of worms, multiplication of the parasite was enhanced by immunosuppression of the host.

MATERIALS AND METHODS

S~rongyloides skxorulis was derived originally from infected humans (Grove, 1980) and since that time has been passaged through dogs, as described previously (Grove & Northern, 1982). Furthermore, persistent and disseminated infections result in dogs immunosuppressed with pred- nisolone, as described earlier (Grove, Heenan & Northern, 1983).

A male, mongrel dog, I 5 kg in weight, was obtained from a pound then was infected with 5000 infective larvae. Four weeks later, the administration of prednisolone 50 mg per day was begun. Larvae were excreted in the faeces over the next 9 months, with counts ranging between 50 and 300 larvae/g faeces. The immunosuppressive dose was then doubled, and 1 month later, the numbers of larvae excreted in the faeces has risen to 22,000/g. The dog was then prepared for examination. The method of in viva fixation of the tissue was similar to that described by Thompson, Dunsmore & Hayton (1979). Briefly, the small intestine was brought to the exterior while the dog was anaesthetised with halothane. A segment of small bowel 10 cm in length and located 20 cm below the pyloric sphincter was ligated but the mesentric blood supply was left intact. Paraformaldehyde-glutar- aldehyde fixative (Graham & Karnovsky, 1966) was injected into the lumen of the ligated segment until it was distended fully. The dogwas then killed, while still under anaesthesia. by intravenous injection of an overdose of pentobarbitone sodium.

The blood vessels were then ligated and the segment was excised. The removed small intestine was cut into rings several millimetres in width and immersed in 4% formalde- hyde or in 2.5% glutaraldehyde in 0.1 M phosphate buffer pH 7.2 at 4 ‘C. Specimens fixed in formalin were embedded in paraffin and sections 6 pm thick were cut and stained with haematoxylin and eosin. Specimens for electron micro- scopical examination were trimmed to 1.0 mm’, washed in phosphate buffer, postfixed in 1% osmium tetroxide in 0.1 M phosphate buffer (pH 7.2) for 1 h, then rewashed in phos- phate buffer. Specimens were then dehydrated in graded solutions of ethanol and propylene oxide and embedded in araldite. Sections 50 nm thick were cut on an LKB ulta- microtome (LKB, Sweden), stained with lead citrate, then examined in Phillips 301 and 410 electron microscopes at an accelerating voltage of 80 kV.

RESULTS

Vast numbers of adult worms were present and were easily seen at both the light and electron micro- scopical levels. The adult worms were observed in close proximity to the epithelial cells. Histological sections disclosed worms deep in the crypts of Lieberkuhn (Fig. 1). Many worms were located within vacuoles adjacent to the lumen and which appeared to be formed by a thin rim of distorted and distended epithelial cells (Fig. 2), although portions of some

worms were noted free in the intestinal lumen. Serial sections indicated that these vacuoles were really tunnels in which the worms lay.

Greater detail of the interface between worm and host tissue was provided by electron microscopical examination. Figures 3 and 4 show complete cross- sections of adult worms at two different levels along the length of the parasite. The worms lay in tunnels between a number of enterocytes which were com- pressed and distorted. There was no evidence of syncytial formation but occasionally the epithelial cells were necrotic (Figs. 5-6). The microvillous border of some of the enterocytes could be clearly seen (Figs. 3-6). Adult worms were never observed deep to the basement lamina. A sparse infiltrate of plasma cells and lymphocytes was noted in the vicinity of the worms (Figs. 6-7). Examination of the interface between the worms and the enterocytes revealed an electron-dense fibrillogranular deposit on the surface of the enterocytes forming the tunnels (Fig. 8). The deposit often had projections which coincided with grooves on the external cortical layer of the worm cuticle (Fig. 9). Furthermore, deposits of this material were also seen more distant from the parasite in the intercellular spaces between the enterocytes (Fig. 8). High-power examination of the interface indicated that the integrity of the enterocyte plasma membrane adjacent to the worm cuticle was sometimes broken (Fig. 10). Eggs/larvae were never observed, either in the lamina propria or in the tunnels between the epithelial ceils.

DISCUSSION

Knowledge of the anatomical location of S. ster- coralis in the small bowel is essential for an under- standing of the mechanisms of both immune expulsion of helminths from the gut or evasion of immune responses leading to persistence of worms in the gut. Furthermore, it may shed some light on the mechan- ism by which the unusual phenomenon of auto- infection occurs in this infection as compared with most other helminthiases, including infections with other species of Strongyloides such as S. ratti. This study has clearly shown that S. stercoralk does penetrate the intestinal mucosa with female adult worms being located within the glandular epithelium, but we found no evidence of either adult worms or larvae in the tissues deep to the basement lamina.

Histological examination of serial sections suggested that S. stercoralis wound its way through the mucosa.

Adult worms appeared to be in tunnels situated in close proximity to the columnar epithelium. Occasionally, portions of adult worms were seen in the lumen of the intestine. Like S. ratti, S. stercoralis appears on the light microscopical level to occupy a niche very similar to that held by Trichinella spiralis. Gardiner (1976) concluded that the adult forms of the latter parasite, which were also found mostly at the bases of thevilli and in the crypts, were embedded in the epithelial layer of the mucosa, while Despommier, Sukhdeo & Meero-

Intestinal location of Strongyloides stercoralis

FIG.

FIG.

1. Low power light microscopical view of small intestine showing cross-sections of adult worms (w) in the crypts of Lieberkuhn and at the bases of the villi. Scale bar = 100 ,U m.

2. High power light microscopical view of cross-sections of adult worms in the crypts of Lieberkuhn. Note the distended epithelial lining (e) and the vacuole(v) around the worm. Scale bar = 40 y m.

vitch (1976) thought that adult Trichinellu spiralis were located beneath the epithelium and above the lamina propria. Furthermore, both papers reported that larvae were released directly into the lamina propria by adult worms.

More recently, Wright (1979) examined the intes- tinal location of T spiralis by electron microscopy. He believed that the adult worms were intracellular rather than extracellular in location and occupied both absorptive and goblet epithelial cells. Although our electron micrographs are very similar to those taken by Wright (1979) of T. spiralis, we believe that

like S. ratti, S. stercoralis adult worms lie between distorted, compressed and displaced columnar epithelial cells, with several enterocytes surrounding each worm. lnterdigitations were noted between the epithelial cells surrounding worms, thus confirming their separate identity. Moreover, it would seem unlikely that enterocytes could survive the continuous plasmalemmal disruption and direct continuity with the extracellular space that intracytoplasmic invasion would require. Nevertheless, the plasma membrane of enterocytes adjacent to a worm was sometimes disrupted, and occasionally cells became necrotic.

1260 D. I. GROVE, A. WAW+ON, L. L. Yu, C. NORTHERN and J. M. PAFADIMITRIOU

FIG. 3. Transmission electron micrograph of an adult worm (w) showing the surrounding vacuole (v) and the enterocyte! forming the tunnel. Note the microvilli (mv) of the enterocytes and the small bowel lumen (L). Scale bar = 6 fi m.

; W

FIG. 4. Transmission electron micrograph of adult worms (w) showing a goblet cell (g) in close proximity to the enterocytes (e). Scale bar = 6 pm.

Intestinal location of S~rongyloides slercorab 1261

FIG. 5. Transmission electron micrograph of an adult worm (w) showing necrotic cells (n) and amorphous material (a). SC bar=6pm.

,ale

FIG. 6. Transmission electron micrograph of an adult worm showing the margin of a necrosed enterocyte (n) and a plasma cell (p) adjacent to it. w = worm. Scale bar = 6 y m.

D. I. GROVE, A. WARTON, L. L. Yu, C. NORTHERN and J. M. PAPADIMITRIOU

7. Transmission electron micrograph showing an adult worm surrounded by inflammatory cells. p = plasma cell, L lymphocyte, w = worm. Scale bar = S ,u m.

8. High power transmission electron micrograph showing an electron-dense deposit (d) on the inner margin ol vacuole and between the enterocytes (e), w = worm. Scale bar = I.2 ,u m.

,v =

the

263

FIG.

FIG.

9. High power transmission electron micrograph showing the electron-dense deposit (d) on the inner margin of enterocytes (e). Note the projection (pr) opposite the annulations in the cuticle (c)of the worm. Scale bar = 0.7 pm.

10. High power transmission electron micrograph showing disruption (arrow) of the cell membrane (cm) of enterocyte (e) in close proximity to the cuticle (c)of the worm. Scale bar = 0.5 pm.

the

the

1264 D. I. GROVE, A. WARTON, L. L. Yu, C. NORTHERN and J. M. PAPADIMITRIOU

These changes may be a consequence of enzymes released by the worms. A fibrillar and granular electron-dense deposit was seen on the surface of the enterocytes opposed to the worms, and this material often infiltrated into the spaces between the entero- cytes. The nature of this deposit is uncertain, but may be derived from helminth, host, or both. Possibilities include antigen-antibody complexes and the rem- nants of necrosed cells. Since the worms are located in the intestinal mucosa between epithelial cells, they are open to attack by inflammatory cells. Because of the profound immunosuppression of the host in this experiment, however, we noted only a sparse inflam- matory infiltrate.

It seems probable that an invading worm is able to fracture junctional complexes between adjoining enterocytes and move actively between them, forming (in cross-section) an apparent vacuole around the worm. This vacuole could be either real or an artefact. Some evidence in favour of the latter is perhaps provided by Figure 9 which shows projections of the fibrillo-granular deposit on the surface of enterocytes opposite depressions in the external cortical layer of the worm cuticle, yet the two are no longer contiguous. On the other hand, observations with S. ratti have shown that the vacuole was not present at the cephalic end of the worm, but that the vacuole became obvious as sections were cut more caudally (Dawkins et al., 1983). We did not observe such a phenomenon in this study, but that may simply represent a sampling error. It seems reasonable to assume that the tunnel rep- resents a fluid-filled space bathing the worm, resulting from the movement of the parasite through the epithelium.

In contrast to T. spiralis in which newborn larvae are found in the lamina propria and deeper parts of the small intestinal wall, we never observed rhab- ditiform or filariform larvae of S. stercoralis in this location. It seems unlikely that this is a sampling error as a large number of sections were examined and multiple adult worms were seen. Likewise, larvae were never observed in the tunnels and it seems probable that embryos are released directly into the small intestinal lumen either when adult worms wander through the lumen or when the vulvar region of the female worms is exposed to the small bowel lumen. Alternatively, rhabditiform larvae could be deposited in the tunnels then quickly move through them into the intestinal lumen. These findings suggest that it is unlikely that autoinfection occurs by rhabditiform larvae being deposited directly in the mucosa and migrating through the tissues. They provide indirect support for the hypothesis that rhabditiform larvae are released into the small intestinal contents, moult twice, then infective larvae penetrate the mucosa lower down in the intestinal tract.

In conclusion, it appears that S. stercoralis adult worms move through the epithelium creating tunnels lined by enterocytes. It seems probable that they exit into the lumen and re-enter the epithelial layer. Adult

worms do not traverse the basement lamina and probably deposit eggs/larvae directly into the small intestine lumen. These observations are consistent with the view that autoinfection results from rhab- ditiform larvae moulting within the lumen of the gastrointestinal tract then filariform larvae penetrat- ing the colonic mucosa.

Acknowleu’gements-This study was supported by a grant from the National Health and Medical Research Council of Australia.

REFERENCES

ABADIE S. H. 1963. The life cycle of Sfrongyloides raf/i. Journal OfParasitology 49: 241-248.

ASKANAZY M. 1900. Ueber Art und Zweck der Invasion der Anguillula intestinalis in die Darmwand. Centralblattfiir Bakteriologie, Parasitenkunde und Infektionskrankhei- ten, Abteilung Originale 27: 569-578. Translated in: Tropical Medicine and Parasitology: Classic Investigations (Edited by KEAN B. H., MOTT K. E. & RUSSELL A. J.), two volumes, pp. 677. Cornell University Press, Ithaca, 1978.

BELDING D. L. 1942. Textbook of ClinicalParasitology, 2nd edn., pp. 888. Appleton Century, New York.

DAWKINS H. J. S., ROBER-~SON T. A.. PA~AUIMITKIOU J. M. & GROW D. I. 1983. Light and electron micro- scopical studies of the location of Strongyloides ratti in the mouse intestine. Zeitschrift fiir Parasitenkunde. Para- sitology Research 69: 357-370.

DESPOMMIER D. D., SUKHDEO M. & MEEROVITCH E. 1976. Trichinefla spiralis: site selection during the enteral phase of infection in mice. Experimental Parasitology 44: 209- __ 215.

FAUST E. C., RUSSELL P. F. & JUNG R. C. 1970. Craig and FaustS Clinical Parasitoloff/, 8th edn.. oo. 890. Lea and Febiger, Philadelphia. __

. .

CARDINER C. H. 1976. Habitat and reoroductive behaviour of Trichinella spiralis. Journal of P’arasitology 62: 865- 870.

GENTA R. M. & WARD P. A. 1980. The histopathology of experimental strongyloidiasis. American Journal 01 Pathology 99: 207-220.

GILLES H. M. 1984. Intestinal nematode infections. In: Hunter’s Tropical Medicine (Edited by STRICKLAND

G. T.), 6th edn., pp. 620-647. GOLGI C. & MONTI A. 1885. Sulla storia naturale e sul

significato clinico-patologica delle cosi-dette anguillule stercorali e intestinali. Atti della Reale Accademia delle Scienze di Torino 21: 55-59.

GRAHAM R. & KARNOVSKV M. J. 1966. The early stages of absorption of injected horseradish peroxidase in proximal tubules of mouse kidney. Journal of Histochemistry and Cytochemistiy 14: 291-302.

GROVE D. I. 1980. Strongyloidiasis in Allied ex-prisoners of war in Southeast Asia. British MedicalJournal 280: 59% 601.

GROVE D. I., HEENAN P. J. & NORTHCRN C. 1983. Persistent and disseminated infections with Strongyloides stercoralis in immunosuppressed dogs. International Journal for Parasitology 13: 483-490.

GROVE D. I. & NORTHERN C. 1982. Infection and immunitv in dogs infected with a human strain of Strongyloides stercoralis. Transuctions of the Royal Society of Tropical Medicine and Hygiene 76: 833-838.

Intestinal location of Strongyloides stercoralis 1265

MAEGRAITH B. G. 1984. Adams and Maegraith’s Clinical Tropical Medicine. 7th edn. Blackwell Scientific Public- ations, Oxford.

MANSON-BAHR P. E. C. & APTED F. I. C. 1982. Munson’s Tropical Diseases, 18th edn. Bailliere Tindall, London.

MARKELL E. K. 1985. Intestinal nematode infections. In: Pediatric Clinics of North America (Edited by SEIDEL J. S.), Vol. 32, pp. 97 l-986. W. B. Saunders,Philadelphia.

NORMAND L. 1876. Sur la maladie dite diarrhee de Cochin- chine. Comptes Rendus Hebdomadaires des Seances de I’Academie des Sciences 83: 3 16-3 18. nale e rhabdonemiasis. Rivisfa Generale ltaliana di Clinica Medka, Pisa 3: 47-56.

THOMPSON R.C.A., DUNSMORE J.D.& HAYTON A.R. 1979. Echinococcus granulosus: secretory activity of the rostellum of the adult cestode in situ in the dog. Experi- mental Parasitology 48: 144-163.

WERTHEIM G. & LENCY J. 1965. Growth and development of Strongyloides ratti Sandground, 1925, in the albino rat. Journal ofParasitology 51: 636-639.

WRIGHT K. A. 1979. Trichinella spiralis: an intracellular parasite in the intestinal phase. JournalofParasitotogv 65: 441-445.