In hyper-infection syndrome, complete disruption of the GI mucosa, ulcerations, paralytic ileus with exudative enteropathy as well as massive GI bleeding may also occur due to the direct invasion of the larvae. Profound diarrhea, malabsorption with consequent hypo-albuminemia and electrolyte disturbances were all consistent with hyper-infection related enteropathy in our patient. On the other hand, effective anthelminthic treatment in hyper-infected patients can lead to mass-destruction of intraluminal and intramural larvae and to release of huge amounts of different toxic inflammatory and vaso-active compounds[7,8].
- Transmission of Strongyloides stercoralis Through Transplantation of Solid Organs — Pennsylvania, 2012 (April 12, 2013 / Vol. 62 / No. 14)
What is added by this report?
Donor-derived Strongyloides infection might be more common than previously believed. In these investigations, a single donor was the source of infection for three of four organ recipients. Testing of pretransplant serum contributed to the determination that infection was donor derived.
Strongyloides stercoralis is an intestinal nematode endemic in the tropics and subtropics. Immunocompetent hosts typically are asymptomatic, despite chronic Strongyloides infection. In contrast, immunocompromised patients are at risk for hyperinfection syndrome and disseminated disease, with a fatality rate >50% (1–3). The infection source for immunocompromised patients, such as solid organ transplant recipients, is not always apparent and might result from reactivation of chronic infection after initiation of immunosuppressive therapy or transmission from the donor. In October 2012, the United Network for Organ Sharing (UNOS) notified CDC of a left kidney and pancreas recipient in Pennsylvania diagnosed with strongyloidiasis. This report summarizes the results of the investigation of the source of Strongyloides infection in three of four organ recipients. Testing of pretransplant donor and recipient sera confirmed that infection in the recipients was donor derived. This investigation underscores the importance of prompt communication between organ procurement organizations, transplant centers, and public health authorities to prevent adverse events in recipients when transmission is suspected. Additionally, it emphasizes the utility of stored pretransplant samples for investigation of suspected transplant-transmitted infections and the need to consider the risk for Strongyloides infection in organ donors.
On October 4, 2012, UNOS notified CDC of a left kidney and pancreas transplant recipient diagnosed with strongyloidiasis. UNOS also identified three additional organ recipients: the right kidney recipient, who received his transplant at the same institution as the index case; the liver recipient, who died within a few days after the transplantation; and the heart recipient, who was diagnosed with suspected reactivation of chronic strongyloidiasis 2 weeks earlier. CDC requested stored pretransplant serum from all organ recipients, along with stored donor serum for testing, to determine if infection with Strongyloides in the recipients was donor derived or reactivation of chronic infection. Evaluation of these specimens revealed that no recipient had detectable Strongyloidesantibody before transplantation, but the donor had evidence of chronic infection based on positive serologic results.
Organ donor. In July 2012, a Puerto Rico-born Hispanic man, aged 24 years, was admitted to a local emergency department with multiple gunshot wounds. After a 9-day hospitalization, he died, and his heart, kidneys, pancreas, and liver were transplanted into four recipients the next day. History obtained from his mother indicated that the donor was a healthy young male who often visited Puerto Rico. Strongyloides infection risk was not considered; therefore, testing was not performed before organ recovery.
Kidney and pancreas recipient. This recipient is a U.S.-born white man, aged 64 years, with end-stage renal disease secondary to long-standing diabetes mellitus who had never traveled outside the United States. Nine weeks posttransplant, he developed severe nausea, anorexia, and abdominal distention and was admitted to the hospital. Stool studies and biopsies performed during an esophagogastroduodenoscopy revealed S. stercoralis adult worms; larvae were found in urine studies. The patient was treated with ivermectin and albendazole, and after a hospitalization complicated by Enterobacter cloacae bacteremia, periduodenal abscess, and loss of pancreatic transplant function, he was discharged in stable condition on ivermectin. Repeat stool analyses were negative 3 days after starting therapy.
Kidney recipient. This recipient is a U.S.-born adolescent, aged 14 years, with end-stage renal disease as a result of a single dysplastic kidney; he had never traveled outside the United States. He was contacted for evaluation 10 weeks posttransplant, after the left kidney and pancreas recipient received a diagnosis of strongyloidiasis. He was discovered to be ill with fever, rash, malaise, anorexia, nausea, vomiting, and diarrhea. He was diagnosed with strongyloidiasis via esophagogastroduodenoscopy-obtained biopsy and stool testing. He was treated with ivermectin for 4 weeks and albendazole for 2 weeks. Repeat stool specimens were negative 3 days after starting therapy and remained negative as of November 2012.
Liver recipient. This recipient was a Hispanic man, aged 66 years, with a history of hepatic failure secondary to chronic hepatitis C infection. He tolerated surgery and was clinically stable until postoperative day 4, when his heart stopped and he was unresponsive to attempts at resuscitation. At autopsy, no evidence of Strongyloides infection was found; cause of death was undetermined.
Heart recipient. This recipient was a U.S.-born Hispanic man, aged 59 years, with ischemic cardiomyopathy; he lived in Puerto Rico for 6 months as a teenager. He remained clinically stable posttransplant and was discharged 11 days after surgery. He experienced multiple episodes of organ rejection and was treated with high doses of steroids. Seven weeks posttransplant, he was readmitted to the hospital with fever and a respiratory illness and required intubation in response to rapid decompensation. He was diagnosed with a viral respiratory illness and given oseltamivir and antibiotic and antifungal medications. A bronchoscopy performed on hospital day 3 showed S. stercoralis larvae. He was started on ivermectin and albendazole for treatment of suspected reactivated chronic strongyloidiasis. He developed gram-negative and enterococcal bacteremia and vancomycin-resistant enterococcal meningitis and became neurologically compromised. Life support was withdrawn, and he died 11 weeks posttransplant.
Anjum Hasan, MD, Marie Le, MD, Jessica Pasko, MD, Karen A. Ravin, MD, Geisinger Medical Center; Heather Clauss, MD, Temple Univ Hospital; Richard Hasz, MFS, Gift of Life Donor Program; Elizabeth A. Hunt, MPH, Pennsylvania Dept of Health. Elizabeth Bosserman, MPH, Isabel McAuliffe, MS, Susan P. Montgomery, DVM, Div of Parasitic Diseases and Malaria, Center for Global Health; Matthew J. Kuehnert, MD, Susan N. Hocevar, MD, Div of Healthcare Quality Promotion, National Center for Emerging and Zoonotic Infectious Diseases; Francisca Abanyie, MD, EIS Officer, CDC. Corresponding contributor: Francisca Abanyie, email@example.com, 404-718-4775.
Most Strongyloides infections in organ transplant recipients are thought to be caused by reactivation of chronic infection after initiation of immunosuppressive therapy. Donor-derived infection has been reported, but the incidence of transmission is unknown (4,5). During 2009–2012, CDC assisted in seven investigations of organ donors and associated recipients with strongyloidiasis determined to be donor derived. Donor-derived infection is difficult to prove, especially if the infected recipient is from a region in which Strongyloides is endemic. Archived pretransplant serum samples were available for recipient testing in this investigation. Results of that testing contributed to the determination that infection was donor derived and not reactivated chronic infection in the recipients.
This investigation revealed several gaps in current understanding and assessment of the risk for transplant-transmitted strongyloidiasis. Specific recommendations are lacking for Strongyloides testing of organ donors from areas in which it is endemic. The parasitic infections sections of the American Society for Transplantation’s guidelines for screening prior to solid organ transplantation recommend testing donors and recipients for Toxoplasma and Trypanosoma cruzi (the cause of Chagas disease), but only recommend screening for Strongyloides in recipients from areas in which the nematodes are endemic, with no mention of donor screening (6,7). These guidelines are not policy, thus screening of donors and recipients for parasitic infections is voluntary, resulting in varied practices among organ procurement organizations and transplant centers based on the perceived risk in their respective patient populations. The growing evidence of transplant transmission of Strongyloides, reported here and in the recent literature, might support development of recommendations for specific testing of donors and recipients from endemic regions to prevent severe strongyloidiasis in recipients (1,4,5). A minimum of three serial stool examinations for larvae, using specialized concentration techniques, is the gold standard for diagnosis of Strongyloides infection, but this might not be feasible in patients who have poor gastrointestinal function or are brain dead. Tests to detect parasite-specific antibody, such as an enzyme-linked immunoassay, also are available and are valuable in identifying Strongyloides infection (8). If infection is confirmed in the donor, prophylaxis could be given to recipients to avert adverse outcomes.
Rapid communication among transplant centers with patients who received organs from a single donor also is essential. The Organ Procurement and Transplant Network encourages organ procurement organizations and transplant programs to communicate promptly through its Patient Safety System, especially when there is concern for potential transmission of disease or medical conditions to the organ recipient from the donor. Such communication ideally should occur within 24 hours after knowledge of or concern for transmission, because multiple recipients might be adversely affected (9).
This investigation illuminates two gaps that need to be filled to improve transplant safety in solid organ recipients at risk for Strongyloides infection: 1) developing recommendations for screening of donors from Strongyloides-endemic areas, and 2) improving communication among transplant centers and organ procurement organizations. Advances in these areas might be life-saving for immunocompromised hosts.
Christine McGarry, Gift of Life Donor Program; Justine Gaspari, Milton S. Hershey Medical Center, Pennsylvania. Patricia Wilkins, PhD, Div of Parasitic Diseases and Malaria, Center for Global Health, CDC.
- Roxby AC, Gottlieb GS, Limaye AP. Strongyloidiasis in transplant patients. Clin Infect Dis 2009;49:1411–23.
- Cappello M, Hotez PJ. Intestinal nemaodes: Strongyloides stercoralis and Strongyloides fuelleborni. In: Long S, Pickering LK, Prober CG, eds. Principles and practice of pediatric infectious diseases. 3rd ed. Philadelphia, PA: Churchill Livingstone-Elsevier; 2008.
- CDC. Parasites—Strongyloides: resources for health professionals. Atlanta, GA: US Department of Health and Human Resources, CDC; 2012. Available at http://www.cdc.gov/parasites/strongyloides/health_professionals/index.html.
- Hamilton KW, Abt PL, Rosenbach MA, et al. Donor-derived Strongyloides stercoralis infections in renal transplant recipients. Transplantation 2011;91:1019–24.
- Weiser JA, Scully BE, Bulman WA, Husain S, Grossman ME. Periumbilical parasitic thumbprint purpura: Strongyloides hyperinfection syndrome acquired from a cadaveric renal transplant. Transpl Infect Dis 2011;13:58–62.
- Fischer SA, Avery RK; AST Infectious Disease Community of Practice. Screening of donor and recipient prior to solid organ transplantation. Am J Transplant 2009;9(Suppl 4):S7–18.
- Anonymous. Screening of donor and recipient prior to solid organ transplantation. Am J Transplantation 2004;4(Suppl 10):10–20.
- Genta RM. Predictive value of an enzyme-linked immunosorbent assay (ELISA) for the serodiagnosis of strongyloidiasis. Am J Clin Path 1988;89:391–4.
- Organ Procurement and Transplantation Network. Identification of transmissible diseases in organ recipients. Rockville, MD: US Department of Health and Human Services, Health Resources and Services Administration, Organ Procurement and Transplantation Network; 2010. Available at http://optn.transplant.hrsa.gov/policiesandbylaws2/policies/pdfs/policy_16.pdf .
Strongyloides stercoralis, a worldwide-distributed soil-transmitted helminth, causes chronic infection which may be life threatening.
Limitations of diagnostic tests and nonspecificity of symptoms have hampered the estimation of the global morbidity due to strongyloidiasis. This work aimed at assessing S. stercoralis-associated morbidity through a systematic review and meta-analysis of the available literature. MEDLINE, Embase, CENTRAL, LILACS, and trial registries (WHO portal) were searched. The study quality was assessed using the Newcastle-Ottawa scale. Odds ratios (ORs) of the association between symptoms and infection status and frequency of infection-associated symptoms were calculated. Six articles from five countries, including 6,014 individuals, were included in the meta-analysis-three were of low quality, one of high quality, and two of very high quality. Abdominal pain (OR 1.74 [CI 1.07-2.94]), diarrhea (OR 1.66 [CI 1.09-2.55]), and urticaria (OR 1.73 [CI 1.22-2.44]) were associated with infection. In 17 eligible studies, these symptoms were reported by a large proportion of the individuals with strongyloidiasis-abdominal pain by 53.1% individuals, diarrhea by 41.6%, and urticaria by 27.8%. After removing the low-quality studies, urticaria remained the only symptom significantly associated with S. stercoralis infection (OR 1.42 [CI 1.24-1.61]). Limitations of evidence included the low number and quality of studies. Our findings especially highlight the appalling knowledge gap about clinical manifestations of this common yet neglected soil-transmitted helminthiasis. Further studies focusing on morbidity and risk factors for dissemination and mortality due to strongyloidiasis are absolutely needed to quantify the burden of S. stercoralis infection and inform public health policies.
…in a Patient with a History of Systemic Lupus Erythematosus
Strongyloidiasis is an infection caused predominantly by the helminth Strongyloides stercoralis. This nematode is endemic to tropical and subtropical regions such as Southeast Asia, but is also present in more temperate climates, such as the northern United States and Canada.1 Infection can rarely occur in areas thought to be non-endemic for the disease. Most chronically infected persons are asymptomatic. Clinical manifestations, when present, are usually mild and non-specific.2
Immunosuppression places infected persons at risk for the Strongyloides hyperinfection syndrome (SHS), where the organism proliferates unchecked. This syndrome can cause exacerbation of the patient’s symptoms related to an increased parasite load in the intestine and lungs. Additional symptoms may arise as the organism involves organs not normally associated with the auto-infective life cycle.2,3 We describe an unusual case of SHS in a patient undergoing chronic corticosteroid treatment for systemic lupus erythematosus (SLE). We review the literature regarding SLS in immunosuppressed patients, with emphasis on those with a history of SLE.
A 30 year-old Hispanic man with an eight-year history of poorly controlled SLE came to an emergency department with fever, diffuse generalized pain, and bilateral upper and lower extremity edema. He was treated with antibiotics and methylprednisolone for presumed sepsis and lupus flare. The patient’s symptoms eventually resolved, but he was found to have nephrotic range protein and erythrocyte casts in his urine. He underwent an ultrasound-guided left renal biopsy, which later confirmed class IV G lupus nephritis. The next day, the patient’s systolic blood pressure decreased to 90 mm of Hg, and he began to experience diffuse abdominal pain, rebound tenderness, guarding, rigidity, and emesis. His leukocyte count and lactate dehydrogenase level were increased, and his hemoglobin level decreased significantly.
Based on the clinical examination and findings of a computed tomographic (CT) angiography of the abdomen and pelvis (Figure 1), the patient underwent an emergent exploratory celiotomy. Blood clots were visualized in the peritoneal cavity, as well as active slow bleeding from the gastrocolic ligament and the base of the transverse mesocolon. Hematomata were identified in the omental bursa, pelvis, and hepatic flexure. No additional source of peritoneal bleeding was identified. The combined operative and CT findings suggested that the vascular supply to the distal transverse colon was compromised. An extended right hemicolectomy with a colonic mucous fistula and end ileostomy was performed.
Grossly, the serosa of the colon was covered by dark red-brown blood but was otherwise unremarkable. Several blood clots were seen within the mesentery and the omentum. The colonic mucosa was diffusely edematous with patches of yellow-tan exudate. There was a mild loss of the mucosal folds with focal edema. No lesion, ulceration, or perforation was identified. Microscopically, there were patchy areas of acute inflammatory cells and cellular debris overlying eroded mucosa. The lamina propria was markedly expanded by a lymphoplasmacytic infiltrate with scattered neutrophils and eosinophils.
There were numerous filariform larvae and sharply pointed, curved tailed adult worms present within luminal acellular debris overlying the ulcerated mucosa. Similar organisms were seen in the lamina propria infiltrating into and running alongside intact crypts (Figure 2). Numerous organisms were seen in the lymphatics (Figure 3).
Treatment of the patient’s Strongyloides hyperinfection was started with a 21-day course of ivermectin and albendazole. The patient then showed development of diffuse alveolar hemorrhage causing acute respiratory distress syndrome. A transbronchial lung biopsy was performed, which showed evidence of cytomegalovirus pneumonia, verified by immunohistochemical stainings. The lung biopsy specimen was remarkable for the presence of a giant cell granulomatous inflammatory response surrounding a filariform larva that presumably died secondary to the anti-helminthic agents (Figure 4). His cytomegalovirus pneumonia was treated with intravenous ganciclovir. His previously mentioned class IV lupus nephritis was treated with intravenous immunoglobulin and pulse steroids with steroid taper.
The resolution of his infection was confirmed with triplicate negative stool ova and parasites studies. Finally, on hospital day 60, he was discharged to a long-term rehabilitation facility. To date, he has no documented sequelae from his infection with Strongyloides.
The interest in this case stems from histopathologic diagnosis of SHS in a patient without any relevant medical history. The patient’s medical history was remarkable only for SLE. The patient indicated a history of professional boxing, which may imply a history of extensive travel; however, this notion is speculative at best. The patient’s condition at admission closely mimicked symptoms described in the patient’s rheumatologic disorder. Accordingly, the index for suspicion for parasitic infections was low to non-existent. Were it not for the series of events that lead to the eventual histopathologic diagnosis, the patient likely would have experienced progression of the hyperinfection syndrome and eventual death.
The life cycle of Strongyloides can either be an isolated free-living cycle where the helminth lives independently in soil, as well as a parasitic cycle in which the infective filariform larvae enter the host via intact skin, mature to adults, and proliferate. The rhabtidiform larvae created in the parasitic cycle are either passed in the stool or re-enter the circulation as filariform larvae by penetrating bowel mucosa or perianal skin to perpetuate the parasitic life cycle. This autoinfection cycle differentiates S. stercoralis from many other helminths,1,4–6 and enables the organism to reside within the host for years, or even decades.
Most persons infected with S. stercoralis are asymptomatic. Clinical manifestations, when present, are often mild and involve the intestine (abdominal pain, diarrhea, constipation, nausea, and weight loss), the skin (rash and pruritus, particularly at the site of entry of the larvae), and the lungs (cough, tracheal irritation, wheezing, and asthma).2,3 The lack of specificity of the clinical syndrome, combined with a lack of sufficiently sensitive diagnostic tests, suggest that the current estimated prevalence of 3–100 million infected persons worldwide1,4 is likely to be a significant underestimate.
Immunosuppressive states place patients at risk for SHS. Although the diagnosis of hyperinfection is not clearly defined, it generally occurs when the immune status of the patient changes, and the organism proliferates unchecked and enters organs not normally involved in the worm’s normal intra-host life cycle. The patient may then show systemic manifestations, or more localized symptoms related to each organ the worm involves (e.g., meningitis or biliary obstruction). Invasion of the larvae through the bowel mucosa may also lead to secondary gram-negative septicemia as gut flora enter the blood stream in tandem with the larvae.2,4,5 The patient may then undergo multi-organ dysfunction, septic shock, and die.
A significant proportion of SHS cases are secondary to immunosuppressive drugs and primary immunodeficiency states, such as genetic disorders and hematologic malignancies. Of these contributing factors, corticosteroids are by far the most common precipitating agent.4,7 The exact mechanism of this is unclear; hypotheses range from modulation of the T cell–mediated immune response to suppression of eosinophilia that normally occurs in response to parasitic infections.3,8 Other hypotheses include a possible stimulatory effect of steroids on the adult female’s ability to produce eggs, or on the larval ability to mature.2 Underlying infection with human T cell lymphotropic virus 1 may also affect the T helper immune response and predispose to disseminated strongyloidiasis, to the point where infection with human T cell lymphotropic virus 1 may be suspected if a patient exhibits sub-optimal response to anti-helminthic treatment.9 Interestingly, an association between acquired imunodeficiency syndrome and an increased risk of SHS has not yet been established. The reason for this finding remains unclear.3,4,8
There is no standard method of diagnosing strongyloidiasis. As in our case, histopathologic diagnosis may be rendered by direct visualization of the larvae or adult worms in biopsy specimens. The filariform larvae can be described histologically as a tubular esophagus measuring 180–380 μm in length with a blunted buccal end and notched tail.1,10 Adult worms are considerably longer and are identified by one anterior esophagus and two posterior reproductive organs.11 In intestinal biopsy specimens, these organisms are found in the crypt epithelium, the lamina propria, and submucosa.6,11 Direct detection can also be made in stool specimens. However, some authors recommend analysis of multiple stool samples because a single stool examination may have sensitivity approximating 30%.12,13 Examination of other specimens, such as sputum, duodenal aspirates, ascitic fluid, pleural fluid, peripheral blood smears, and cerebrospinal fluid, may be performed.4,10 The blood agar plate method is a unique and sensitive diagnostic method in which presence of the organism is confirmed by visualizing tracts of bacterial colonies left in the organism’s wake as it travels across the agar plate’s surface.3Newer modalities to detect Strongyloides-specific antigens have been described, such as polymerase chain reaction, which can simultaneously test for presence of other parasitic infections, but can show false-negative results because of potential polymerase chain reaction inhibitors present within patient samples or by inconsistent shedding of the organism in the feces.14
The diagnosis of strongyloidiasis can also be made by using serologic analysis and identification of antibodies against Strongyloides. The enzyme-linked immunosorbent assay has been described as an effective method of testing because of its practicality, ability for automation, and its ability to detect the presence of antibody or antigen, depending on the assay.14,15Similar testing methods have been described; these include dipstick assays, gelatin particle agglutination, and immediate hypersensitivity skin tests to Strongyloides antigens.3,15 Although reasonably effective, these types of tests are prone to cross-reactivity with other helminthic infections and are incapable of differentiating current from past infections.4,14 Moreover, serologic assays may show false-negative results during acute infections or in immunosuppressed patients.3,4,14,16 Assays or studies that directly detect the organism or its antigens may prove helpful in these cases. Luciferase immunoprecipitation system assays have recently been developed and showed promise in detection of Strongyloides-specific antibodies because of its high sensitivity and specificity, lack of cross-reactivity with other parasitic infections, and ability to monitor changes in antibody titers over time, resulting in an effective method of assessing treatment response.14,16,17
Treatment of uncomplicated cases requires standard treatment with anti-helminth drugs, such as ivermectin or albendazole. However, treatment protocols for SHS have not been well established because of lack of data. Furthermore, whether strongyloidiasis patients will benefit from concurrent reduction in immunosuppressive therapy remains debatable. Reports have generally advocated daily anti-helminthic treatment until stool ova and parasite samples are repeatedly negative for an extended period, often up to two weeks.2,4 Response to anti-helminthic therapy is variable in immunosuppressed patients; accordingly, treatment in these patients depends on the etiology of the patient’s immunosuppression.3
Thirteen other cases of SHS occurring in a patient with a history of SLE have been identified. For most of these cases, the diagnosis was either made too late in the disease course to prevent death,18–22 or after the patient succumbed to disease.8,23–25 In only four of these thirteen cases was the diagnosis made and treatment initiated sufficiently early to provide a favorable clinical outcome for the patient.8,10,26,27 Compounding the issue is that these diagnoses were also reported in non-endemic areas, where index of suspicion is low,7 made possible because of the long periods that S. stercoraliscan reside in a host. The high percentage of asymptomatic chronic strongyloidiasis, its non-specific symptoms and similarity in clinical presentation to entities such as the SLE flare (as in our patient), and its appearance in non-endemic areas contribute to the high mortality rate for SHS.5
To prevent development of SHS and hyperinfection, screening for Strongyloides infection has been advocated for patients with a relevant medical history (such as residence or travel in disease-endemic areas) who are either in a immunosuppressed state or about to undergo immunosuppressive treatment.16 Serologic assays have been advocated as primary screening tests because of their reliability and the high sensitivity described in some assays.14 Although some assays are limited in their inability to differentiate current from past infection, some authors state that because of the persistence of strongyloidiasis, a patient with a compatible history and a positive serologic results may benefit from empiric treatment, such as a 1–2 day course of ivermectin if they are immunosuppressed or about to undergo immunosuppressive treatment.3,16
In our patient, the diagnosis was based on identification of an unusual load of worms and filariform larvae detected during the routine histologic examination of the colectomy specimen. A transbronchial pulmonary biopsy obtained soon thereafter isolated larvae in the alveolar septae. In our case, the rapid initiation of anti-helminthic therapy cleared his strongyloidiasis, verified by repeatedly negative stool ova and parasite samples. Our case underscores the importance of maintaining a baseline index of suspicion of stronygloidiasis in immunocompromised patients because this disease is potentially a fatal infection that can be treated successfully with anti-microbial agents.
- Murray PR, Rosethal KS, Pfaller MA, , eds. 2013. Nematodes. Medical Microbiology. Seventh edition. Philadelphia, PA: W. B. Saunders, 778–795. [Google Scholar]
- Concha R, Harrington W, Jr Rogers AI, , 2005. Intestinal strongyloidiasis: recognition, management, and determinants of outcome. J Clin Gastroenterol 39: 203–211.[Crossref] [Google Scholar]
- Keiser PB, Nutman TB, , 2004. Strongyloides stercoralis in the immunocompromised population. Clin Microbiol Rev 17: 208–217.[Crossref] [Google Scholar]
- Ramanathan R, Nutman T, , 2008. Strongyloides stercoralis infection in the immunocompromised host. Curr Infect Dis Rep10: 105–110.[Crossref] [Google Scholar]
- Lam CS, Tong MK, Chan KM, Siu YP, , 2006. Disseminated strongyloidiasis: a retrospective study of clinical course and outcome. Eur J Clin Microbiol Infect Dis 25: 14–18.[Crossref] [Google Scholar]
- Al Samman M, Haque S, Long JD, , 1999. Strongyloidiasis colitis: a case report and review of the literature. J Clin Gastroenterol 28: 77–80.[Crossref] [Google Scholar]
- Buonfrate D, Requena-Mendez A, Angheben A, Muñoz J, Gobbi F, Van Den Ende J, Bisoffi Z, , 2013. Severe strongyloidiasis: a systematic review of case reports. BMC Infect Dis 13: 78.[Crossref] [Google Scholar]
- Mora CS, Segami MI, Hidalgo JA, , 2006. Strongyloides stercoralis hyperinfection in systemic lupus erythematosus and the antiphospholipid syndrome. Semin Arthritis Rheum 36: 135–143.[Crossref] [Google Scholar]
- Hirata T, Uchima N, Kishimoto K, Zaha O, Kinjo N, Hokama A, Sakugawa H, Kinjo F, Fujita J, , 2006. Impairment of host immune response against Strongyloides stercoralis by human T cell lymphotropic virus type 1 infection. Am J Trop Med Hyg 74: 246–249. [Google Scholar]
- Kothary NN, Muskie JM, Mathur SC, , 1999. Strongyloides stercoralis hyperinfection. Radiographics 19: 1077–1081.[Crossref] [Google Scholar]
- Overstreet K, Chen J, Rodriguez JW, Wiener G, , 2003. Endoscopic and histopathologic findings of Strongyloides stercoralisinfection in a patient with AIDS. Gastrointest Endosc 58: 928–931.[Crossref] [Google Scholar]
- Siddiqui AA, Berk SL, , 2001. Diagnosis of Strongyloides stercoralis infection. Clin Infect Dis 33: 1040–1047.[Crossref][Google Scholar]
- Moghadam KG, Khashayar P, Hashemi M, , 2011. Gastrointestinal strongyloidiasis in immunocompromised patients: a case report. Acta Med Indones 43: 191–194. [Google Scholar]
- Levenhagen MA, Costa-Cruz JM, , 2014. Update on immunologic and molecular diagnosis of human strongyloidiasis. Acta Trop 135: 33–43.[Crossref] [Google Scholar]
- van Doorn HR, Koelewijn R, Hofwegen H, Gilis H, Wetsteyn JC, Wismans PJ, Sarfati C, Vervoort T, van Gool T, , 2007. Use of enzyme-linked immunosorbent assay and dipstick assay for detection of Strongyloides stercoralis infection in humans. J Clin Microbiol 45: 438–442.[Crossref] [Google Scholar]
- Mejia R, Nutman TB, , 2012. Screening, prevention, and treatment for hyperinfection syndrome and disseminated infections caused by Strongyloides stercoralis . Curr Opin Infect Dis 25: 458–463.[Crossref] [Google Scholar]
- Ramanathan R, Burbelo PD, Groot S, Iadarola MJ, Neva FA, Nutman TB, , 2008. A luciferase immunoprecipitation systems assay enhances the sensitivity and specificity of diagnosis of Strongyloides stercoralis infection. J Infect Dis 198: 444–451.[Crossref] [Google Scholar]
- Livneh A, Coman EA, Cho SH, Lipstein-Kresch E, , 1988. Strongyloides stercoralis hyperinfection mimicking systemic lupus erythematosus flare. Arthritis Rheum 31: 930–931.[Crossref] [Google Scholar]
- Hughes R, McGuire G, , 2001. Delayed diagnosis of disseminated strongyloidiasis. Intensive Care Med 27: 310–312.[Crossref] [Google Scholar]
- Setoyama M, Fukumaru S, Takasaki T, Yoshida H, Kanzaki T, , 1997. SLE with death from acute massive pulmonary hemorrhage caused by disseminated strongyloidiasis. Scand J Rheumatol 26: 389–391.[Crossref] [Google Scholar]
- Arsić-Arsenijević V, Dzamić A, Dzamić Z, Milobratović D, Tomić D, , 2005. Fatal Strongyloides stercoralis infection in a young woman with lupus glomerulonephritis. J Nephrol 18: 787–790. [Google Scholar]
- Finkielman JD, Grinberg AR, Paz LA, Plana JL, Benchetrit GA, Nicastro MA, Roncoroni AJ, , 1996. Case report: reactive hemophagocytic syndrome associated with disseminated strongyloidiasis. Am J Med Sci 312: 37–39.[Crossref] [Google Scholar]
- Lemos LB, Qu Z, Laucirica R, Fred HL, , 2003. Hyperinfection syndrome in strongyloidiasis: report of two cases. Ann Diagn Pathol 7: 87–94.[Crossref] [Google Scholar]
- Reiman S, Fisher R, Dodds C, Trinh C, Laucirica R, Whigham CJ, , 2002. Mesenteric arteriographic findings in a patient with Strongyloides stercoralis hyperinfection. J Vasc Interv Radiol 13: 635–638.[Crossref] [Google Scholar]
- Rojo-Marcos G, Cuadros-González J, González-Juárez MJ, Gómez-Ayerbe C, , 2009. Strongyloides stercoralis hyperinfection syndrome in a Colombian patient receiving immunosuppressive treatment [in Spanish]. Enferm Infecc Microbiol Clin 27: 432–434.[Crossref] [Google Scholar]
- Wachter RM, Burke AM, MacGregor RR, , 1984. Strongyloides stercoralis hyperinfection masquerading as cerebral vasculitis. Arch Neurol 41: 1213–1216.[Crossref] [Google Scholar]
- Hayden GM, Atlas SA, , 1995. Strongyloidiasis masquerading as inflammatory bowel disease in a patient with lupus erythematosis: a case report. Conn Med 59: 649–650. [Google Scholar]
~Because we needed to be informed that Strongy is a vulgar bugger. In the meantime, I will keep very tight reigns on my mucous membranes.
~Content Source – Strongylus Vulgaris. Following the fibrin tracks
Pathogenesis due to larval migrations
Strongylus vulgaris is the most pathogenic of the large strongyles because of the prolonged (at least 4 months) and extensive migrations through the mesenteric arterial system and its branches before returning to mature in the cecum and colon. Larval migrations cause damage to the smooth endothelial surfaces of arteries, providing a focus for clot formation. These clots (thrombi) are accompanied by inflammation and a progressive thickening of the arterial walls.
The time sequence of lesions caused by migrating larvae following experimental infections are summarized in the table below and have been gleaned from many reports in the literature. A summary of these reports has been given by Ogbourne and Duncan in a 1985 publication from the Commonwealth Institute of Parasitology entitled “Strongylus vulgaris in the horse: its biology and importance.
Time sequence of lesions caused by infections with Strongylus vulgaris
- 0-48 hours after infection – Mucosal hemorrhages in the ileum, cecum and colon.
- 0-7 days after infection – Inflammation of small intestinal arteries in the submucosa and formation of thrombi along the tracks of migrating larvae. Significant infiltration of neutrophils in the submucosa.
- 8-10 days after infection – Arteritis extends through the muscularis mucosa to the serosa.
- 11-21 days after infection – Arteritis extends along all the branches of the ileo-cecal colic artery (supplying the ileum, the dorsal and ventral colon and the cecum) to the cranial mesenteric artery. Arterial walls become thickened and histological sections show a marked cellular infiltration including neutrophils, macrophages, lymphocytes and plasma cells.
- 3 weeks-4 months after infection – The wall of the cranial mesenteric artery is thickened and fibrous and thrombi are associated with the presence of 4th stage larvae and immature adults. Fibrin tracks in the aorta associated with some larvae migrating beyond the cranial mesenteric artery.
- 4-9 months after infection – In the absence of reinfection, arterial lesions heal. By 9 months after infection, the endothelial lining of affected arteries is smooth again and there are few indications of damage other than histological evidence of fibrosis in arterial walls and the presence of macrophages.
In naturally infected animals, arterial lesions are most commonly seen in the cranial mesenteric artery and its branches. However, lesions have also been found less commonly in other arteries including the abdominal aorta, the renal arteries and the celiac axis.
The walls of the cranial mesenteric and the ileo-ceco-colic arteries are invariably thickened and contain large amounts of thrombus material in which are found S. vulgaris larvae. This lesion is properly called verminous arteritis. The lumen of the cranial mesenteric artery is usually constricted in its diameter due to the thickening of the wall and the presence of thrombi. The lumen of smaller arteries may be entirely occluded.
Some reports in the literature describe aneurysms of the cranial mesenteric artery and its branches. True aneurysms with dilation and thinning of the arterial wall due to a loss of elastic fibres are unusual and may result from penetration of the elastic layer of the arterial wall by larvae.
In horses that have died of an acute clinical syndrome, infarction and necrosis of areas of the intestine are usually found at necropsy. These areas of infarction invariably coincide with occlusions (due to thrombi and emboli) in arteries supplying blood to the affected region(s) of the intestine.
These larvae migrate through the host body such that from 24 hours post infection (p.i.) they are found in the naso-frontal region of the host from where they are, presumably, swallowed to reach the small intestine (Tindall and Wilson, 1988).https://www.ncbi.nlm.nih.gov/books/NBK19795/
A prominent and intriguing subgroup of disseminated strongyloidiasis cases are of former far-east prisoners of war (World War II) who now reside in Europe, Australasia, or North America and present with the disease as much as 50 years after leaving the endemic area. Corticosteroid therapy and/or concurrent cancer treatment are common features of these cases (Gill and Bell, 1979; Gill et al., 2004).
The life-cycle of Strongyloides is both complicated and somewhat tricky to understand, but also one part of the fascination of the genus. This description is primarily based on the life-cycle of S. ratti (a parasite of the rat), which has been extensively studied; some of these details are likely to vary in other species.
Hosts become infected when free-living infective L3s penetrate the skin. Naturally this occurs by the chance coming together of host skin and larvae, though this is facilitated by dispersal and nictation behaviours of the larvae. In the laboratory this comes about by the deliberate application of infective L3s to host skin or by subcutaneous injection (Tindall and Wilson, 1988). These larvae migrate through the host body such that from 24 hours post infection (p.i.) they are found in the naso-frontal region of the host from where they are, presumably, swallowed to reach the small intestine (Tindall and Wilson, 1988). This naso-frontal route of migration has been most thoroughly determined in S. ratti; in other species of Strongyloides (and other genera of skin-penetrating nematodes) migration through the lungs is also thought to be important. During this migration they moult via an L4 stage so that there are adult parasitic female worms present in the gut from approximately 4 days p.i., with reproduction commencing shortly thereafter, detected by the presence of eggs and/or larvae in the faeces (Kimura et al., 1999).
In the host faeces the eggs hatch to release first-stage larvae (L1) (Figure 6). Larvae are either male or female. Male larvae develop via L2-L4 stages into rhabditiform males. Analogously, female larvae can develop into rhabditiform females. Together, this type of development is known as indirect, sexual, or heterogonic development. The free-living adults mate and the female lays eggs that hatch to release L1s that moult via an L2 into infective filariform L3 stages. All the progeny of the free-living adult generation are female. These infective L3 stages are long lived and can persist in the environment until they encounter a suitable host. Their behaviour is to move away from the host faeces in which they have developed, a behaviour likely to enhance their probability of finding a host. In addition, female L1s that hatch from eggs passed in faeces have an alternative fate, to moult via an L2 into infective L3s. This type of development is known as direct, asexual, or homogonic development. Infective L3s that have developed via the direct or indirect route are, apparently, the same.
For most Strongyloides species only one free-living adult generation occurs. However, up to nine (decreasingly fecund) free-living generations have been observed for S. planiceps, though in the same study S. stercoralis had only one free-living adult generation (Yamada et al., 1991). Strongyloides therefore differs from its relative Parastrongyloides which has apparently unlimited successive free-living generations (Grant et al., 2006a).
The free-living adult generation of Strongyloides (and Parastrongyloides) is therefore rather similar to the life-cycle of C. elegans, that is, L1-L4 stages that develop into free-living adults that mate to produce eggs and L1 progeny. However, the major difference is that the progeny of Strongyloides spp. free-living adults develop into infective L3s, the hypothesised analogue of dauer larvae (Hotez et al., 1993).
S. stercoralis can undergo autoinfection, that is, repeated generations of development in the same host individual. Autoinfection appears to be unique to S. stercoralis within the genus (and also essentially unique among other genera of gastrointestinal parasites of vertebrates; only Enterobius spp. and Capillaria spp. also have this phenomenon) and largely accounts for it being a serious pathogen of humans. Autoinfection involves accelerated development by larval progeny of parasitic females such that they develop into infective L3s within the gut, which then penetrate directly into the tissues of the primary host. Thus here, the entire life cycle is completed within a host and there are no stages external to the host. Autoinfection may result in dissemination of L3 through many organs and tissues of the host, as well as the establishment of new parasitic females in the gut. In the absence of treatment, subsequent rounds of autoinfection are possible, resulting in fulminant expansion of parasite populations and multi-organ involvement with potentially fatal consequences for the host (Igra-Siegman et al., 1981). Groups at risk of such so-called disseminated S. stercoralis infections include patients who are immunocompromised as a result of corticosteroid therapy, various neoplasms, or infection with the human T lymphotropic virus-1 (Carvalho and Da Fonseca Porta, 2004; Lim et al., 2004; Buofrate et al., 2013).
A prominent and intriguing subgroup of disseminated strongyloidiasis cases are of former far-east prisoners of war (World War II) who now reside in Europe, Australasia, or North America and present with the disease as much as 50 years after leaving the endemic area. Corticosteroid therapy and/or concurrent cancer treatment are common features of these cases (Gill and Bell, 1979; Gill et al., 2004). The onset of disseminated strongyloidiasis decades after the last possible exposure to the parasite is extremely serious for elderly patients. But, this also makes clear a salient feature of the infection biology of S. stercoralis, namely the capacity to maintain exceedingly chronic infections in hosts. Such hyperchronic infections usually go undiagnosed due to the paucity or absence of larvae in the faeces. Alternative hypotheses for the chronicity of these infections are that either there are dormant larvae in the tissues or senescent non-reproductive female worms in the intestine, with immunosuppression triggering either reactivation of dormant larvae or resumption of egg laying by barren parasitic females. Data from studies conducted over a relatively short time frame (77 days) in experimentally infected dogs favour the latter mechanism involving resumed oviposition by barren female worms (Mansfield et al., 1996). The importance of people with chronic asymptomatic strongyloidiasis as a group at risk of disseminated hyperinfection has recently been emphasised (Caumes and Keystone, 2011).
Hypobiosis or dormancy of Strongyloides L3s may or may not be central to the maintenance of chronic infections, but it is key to another mode of transmission: transmammary transmission. There is evidence of transmammary transmission in S. ratti and S. venezuelensis in rats (Nolan and Katz, 1981; Kawanabe et al., 1988), S. stercoralis in dogs (Shoop et al., 2002), S. fuelleborni kellyi in humans (Ashford et al., 1992), and several species affecting livestock including S. ransomi in swine (Stewart et al., 1976), S. westeri in horses (Lyons, 1994), and S. papillosus in ruminants (Moncol and Grice, 1974). Infective L3s transmitted by the transmammary route presumably arrest their development and migration in the mammary glands, and then re-activate at lactation. Transmammary transmission also occurs in other parasitic nematodes that have a phase of within-host tissue migration during their life cycles, including ascarid roundworms and hookworms (Stone and Smith, 1973; Shoop and Corkum, 1987).
Sweat might act as one of the most probable factors for infection by this skin-penetrating nematode.
The host-finding behavior of Strongyloides stercoralis infective larvae was examined by in vitro agarose assay method. As human body fluid contains 0.85% (ca 0.15 molar) NaCl, various concentrations of sodium chloride, from 0.5M to 0.01M (7 steps), were examined.
Many larvae were attracted at concentrations between 0.5 and 0.05M of sodium chloride. The concentration of 0.05M attracted the most larvae. The concentration of 0.02M of sodium chloride showed greatly reduced larval attraction compared with 0.05M. Therefore, the threshold concentration was determined as 0.05M. Then, 0.05M of chemicals were examined in a further experiment. Chloride compounds (NaCl, KCl, CaCl2, MgCl2) were investigated. These chemicals are components of human body fluids.
Distilled water was used as the control in all experiments. Only sodium chloride attracted the larvae. Next, alkaline compounds were examined [NaOH, KOH, Ca(OH)2, and Mg(OH)2]. Larvae accumulated only at the NaOH site. The results suggested that the Na cation is important for larval attraction. A high pH value did not influence attraction at all. Next, human serum was tested. The human serum used was from normal serum to 1:32 diluted sera by distilled water (7 steps). Hierarchical attraction was seen according to serum concentration.
Next, human sweat was collected from a limited zone of chest skin where only eccrine glands were distributed. Non-diluted sweat attracted the most larvae. Sweat might act as one of the most probable factors for infection by this skin-penetrating nematode.
Few other human parasites are associated with such a diverse spectrum of clinical manifestations as Strongyloides stercoralis, yet the basic biological behavior of this unusually versatile worm, particularly with respect to its ability to cause severe disseminated disease in certain hosts, is poorly understood. The current uncritical acceptance of the theory that cell-mediated immunity controls autoinfection has stifled research in other directions. After reviewing what is and is not known about the parasite’s behavior in its host, this article explores some of the mechanisms that could be involved in the regulation of the parasite population. Taking the provocative viewpoint that the parasite, not the host, is mainly responsible for the maintenance of a balanced relationship between the two, I propose a new theory that corticosteroids may act on the intraintestinal larvae as molting hormones and directly promote the development of disseminated disease.
These references are in PubMed. This may not be the complete list of references from this article.
- Agbo K, Deniau M. Anguillulospermie rebelle au traitement. A propos d’un cas diagnostiqué au Togo. Bull Soc Pathol Exot Filiales. 1987;80(2):271–273. [PubMed] [Google Scholar]
- Akoğlu T, Tuncer I, Erken E, Gürcay A, Ozer FL, Ozcan K. Parasitic arthritis induced by Strongyloides stercoralis. Ann Rheum Dis. 1984 Jun;43(3):523–525. [PMC free article][PubMed] [Google Scholar]
- ALCORN MO, Jr, KOTCHER E. Secondary malabsorption syndrome produced by chronic strongyloidiasis. South Med J. 1961 Feb;54:193–197. [PubMed] [Google Scholar]
- Anderson RM. The population dynamics and epidemiology of intestinal nematode infections. Trans R Soc Trop Med Hyg. 1986;80(5):686–696. [PubMed] [Google Scholar]
- ARTHUR RP, SHELLEY WB. Larva currens; a distinctive variant of cutaneous larva migrans due to Strongyloides stercoralis. AMA Arch Derm. 1958 Aug;78(2):186–190. [PubMed] [Google Scholar]
- Badaró R, Carvalho EM, Santos RB, Gam AA, Genta RM. Parasite-specific humoral responses in different clinical forms of strongyloidiasis. Trans R Soc Trop Med Hyg. 1987;81(1):149–150.[PubMed] [Google Scholar]
- Baird JK, De Vinatea ML, Macher AM, Sierra JA, Lasala G. AIDS. Case for diagnosis series, 1987. Mil Med. 1987 Apr;152(4):M17–M24. [PubMed] [Google Scholar]
- Berry AJ, Long EG, Smith JH, Gourley WK, Fine DP. Chronic relapsing colitis due to Strongyloides stercoralis. Am J Trop Med Hyg. 1983 Nov;32(6):1289–1293. [PubMed] [Google Scholar]
- Bhatt BD, Cappell MS, Smilow PC, Das KM. Recurrent massive upper gastrointestinal hemorrhage due to Strongyloides stercoralis infection. Am J Gastroenterol. 1990 Aug;85(8):1034–1036. [PubMed] [Google Scholar]
- Bout D, Haque A, Capron A. Filaricidal effects of cyclosporin-A against Dipetalonema viteae in Mastomys natalensis. Trans R Soc Trop Med Hyg. 1984;78(5):670–671. [PubMed] [Google Scholar]
- Bueding E, Hawkins J, Cha YN. Antischistosomal effects of cyclosporin A. Agents Actions. 1981 Jul;11(4):380–383. [PubMed] [Google Scholar]
- Burke JA. Strongyloidiasis in childhood. Am J Dis Child. 1978 Nov;132(11):1130–1136.[PubMed] [Google Scholar]
- Carp NZ, Nejman JH, Kelly JJ. Strongyloidiasis. An unusual cause of colonic pseudopolyposis and gastrointestinal bleeding. Surg Endosc. 1987;1(3):175–177. [PubMed] [Google Scholar]
- Carvalho Filho E. Strongyloidiasis. Clin Gastroenterol. 1978 Jan;7(1):179–200. [PubMed] [Google Scholar]
- Colebunders R, Lusakumuni K, Nelson AM, Gigase P, Lebughe I, van Marck E, Kapita B, Francis H, Salaun JJ, Quinn TC, et al. Persistent diarrhoea in Zairian AIDS patients: an endoscopic and histological study. Gut. 1988 Dec;29(12):1687–1691. [PMC free article][PubMed] [Google Scholar]
- Corsini AC. Strongyloidiasis and chronic urticaria. Postgrad Med J. 1982 Apr;58(678):247–248.[PMC free article] [PubMed] [Google Scholar]
- Cruz T, Reboucas G, Rocha H. Fatal strongyloidiasis in patients receiving corticosteroids. N Engl J Med. 1966 Nov 17;275(20):1093–1096. [PubMed] [Google Scholar]
- Cummins RO, Suratt PM, Horwitz DA. Disseminated Strongyloides stercoralis infection. Association with ectopic ACTH syndrome and depressed cell-mediated immunity. Arch Intern Med. 1978 Jun;138(6):1005–1006. [PubMed] [Google Scholar]
- Date A, Vaska K, Vaska PH, Pandey AP, Kirubakaran MG, Shastry JC. Terminal infections in renal transplant patients in a tropical environment. Nephron. 1982;32(3):253–257. [PubMed] [Google Scholar]
- Davidson RA. Strongyloidiasis: a presentation of 63 cases. N C Med J. 1982 Jan;43(1):23–25.[PubMed] [Google Scholar]
- Debussche X, Toublanc M, Camillieri JP, Assan R. Overwhelming strongyloidiasis in a diabetic patient following ACTH treatment and keto-acidosis. Diabete Metab. 1988 May-Jun;14(3):294–298. [PubMed] [Google Scholar]
- Dees A, Batenburg PL, Umar HM, Menon RS, Verweij J. Strongyloides stercoralis associated with a bleeding gastric ulcer. Gut. 1990 Dec;31(12):1414–1415. [PMC free article] [PubMed] [Google Scholar]
- de Messias IT, Telles FQ, Boaretti AC, Sliva S, Guimarres LM, Genta RM. Clinical, immunological and epidemiological aspects of strongyloidiasis in an endemic area of Brazil. Allergol Immunopathol (Madr) 1987 Jan-Feb;15(1):37–41. [PubMed] [Google Scholar]
- Brandt de Oliveira R, Voltarelli JC, Meneghelli UG. Severe strongyloidiasis associated with hypogammaglobulinaemia. Parasite Immunol. 1981 Summer;3(2):165–169. [PubMed] [Google Scholar]
- de PAOLA, DIAS LB, da SILVA J. Enteritis due to Strongyloides stercoralis. A report of 5 fatal cases. Am J Dig Dis. 1962 Dec;7:1086–1098. [PubMed] [Google Scholar]
- DeVault GA, Jr, Brown ST, 3rd, Montoya SF, Jr, King JW, Rohr MS, McDonald JC. Disseminated strongyloidiasis complicating acute renal allograft rejection. Prolonged thiabendazole administration and successful retransplantation. Transplantation. 1982 Oct;34(4):220–221. [PubMed] [Google Scholar]
- DeVault GA, Jr, King JW, Rohr MS, Landreneau MD, Brown ST, 3rd, McDonald JC. Opportunistic infections with Strongyloides stercoralis in renal transplantation. Rev Infect Dis. 1990 Jul-Aug;12(4):653–671. [PubMed] [Google Scholar]
- Douce RW, Brown AE, Khamboonruang C, Walzer PD, Genta RM. Seroepidemiology of strongyloidiasis in a Thai village. Int J Parasitol. 1987 Oct;17(7):1343–1348. [PubMed] [Google Scholar]
- Dutcher JP, Marcus SL, Tanowitz HB, Wittner M, Fuks JZ, Wiernik PH. Disseminated strongyloidiasis with central nervous system involvement diagnosed antemortem in a patient with acquired immunodeficiency syndrome and Burkitts lymphoma. Cancer. 1990 Dec 1;66(11):2417–2420. [PubMed] [Google Scholar]
- Fleming AF. Opportunistic infections in AIDS in developed and developing countries. Trans R Soc Trop Med Hyg. 1990;84 (Suppl 1):1–6. [PubMed] [Google Scholar]
- Ford J, Reiss-Levy E, Clark E, Dyson AJ, Schonell M. Pulmonary strongyloidiasis and lung abscess. Chest. 1981 Feb;79(2):239–240. [PubMed] [Google Scholar]
- Forzy G, Dhondt JL, Leloire O, Shayeb J, Vincent G. Reactive arthritis and Strongyloides. JAMA. 1988 May 6;259(17):2546–2547. [PubMed] [Google Scholar]
- GALLIARD H. Recherches sur l’infestation expérimentale a Strongyloides stercoralis au Tonkin (1re note). Ann Parasitol Hum Comp. 1950;25(5-6):441–contd. [PubMed] [Google Scholar]
- GALLIARD H, BERDONNEAU R. Strongyloidose expérimentale chez le chien; effets de la cortisone, résultats du test de Thorn à l’hormone corticotrope (ACTH). Ann Parasitol Hum Comp. 1953;28(3):163–171. [PubMed] [Google Scholar]
- Garcia FT, Sessions JT, Strum WB, Schweistris E, Tripathy K, Bolaños O, Lotero H, Duque E, Ramelli D, Mayoral LG. Intestinal function and morphology in strongyloidiasis. Am J Trop Med Hyg. 1977 Sep;26(5 Pt 1):859–865. [PubMed] [Google Scholar]
- Genta RM. Strongyloides stercoralis: immunobiological considerations on an unusual worm. Parasitol Today. 1986 Sep;2(9):241–246. [PubMed] [Google Scholar]
- Genta RM. Global prevalence of strongyloidiasis: critical review with epidemiologic insights into the prevention of disseminated disease. Rev Infect Dis. 1989 Sep-Oct;11(5):755–767.[PubMed] [Google Scholar]
- Genta RM, Douce RW, Walzer PD. Diagnostic implications of parasite-specific immune responses in immunocompromised patients with strongyloidiasis. J Clin Microbiol. 1986 Jun;23(6):1099–1103. [PMC free article] [PubMed] [Google Scholar]
- Genta RM, Frei DF, Linke MJ. Demonstration and partial characterization of parasite-specific immunoglobulin A responses in human strongyloidiasis. J Clin Microbiol. 1987 Aug;25(8):1505–1510. [PMC free article] [PubMed] [Google Scholar]
- Genta RM, Gatti S, Linke MJ, Cevini C, Scaglia M. Endemic strongyloidiasis in northern Italy: clinical and immunological aspects. Q J Med. 1988 Sep;68(257):679–690. [PubMed] [Google Scholar]
- Genta RM, Harper JS, 3rd, Gam AA, London WI, Neva FA. Experimental disseminated strongyloidiasis in Erythrocebus patas. II. Immunology. Am J Trop Med Hyg. 1984 May;33(3):444–450. [PubMed] [Google Scholar]
- Genta RM, Lillibridge JP. Prominence of IgG4 antibodies in the human responses to Strongyloides stercoralis infection. J Infect Dis. 1989 Oct;160(4):692–699. [PubMed] [Google Scholar]
- Genta RM, Miles P, Fields K. Opportunistic Strongyloides stercoralis infection in lymphoma patients. Report of a case and review of the literature. Cancer. 1989 Apr 1;63(7):1407–1411.[PubMed] [Google Scholar]
- Genta RM, Ottesen EA, Neva FA, Walzer PD, Tanowitz HB, Wittner M. Cellular responses in human strongyloidiasis. Am J Trop Med Hyg. 1983 Sep;32(5):990–994. [PubMed] [Google Scholar]
- Genta RM, Ottesen EA, Poindexter R, Gam AA, Neva FA, Tanowitz HB, Wittner M. Specific allergic sensitization to Strongyloides antigens in human strongyloidiasis. Lab Invest. 1983 May;48(5):633–638. [PubMed] [Google Scholar]
- Genta RM, Schad GA, Hellman ME. Strongyloides stercoralis: parasitological, immunological and pathological observations in immunosuppressed dogs. Trans R Soc Trop Med Hyg. 1986;80(1):34–41. [PubMed] [Google Scholar]
- Genta RM, Weil GJ. Antibodies to Strongyloides stercoralis larval surface antigens in chronic strongyloidiasis. Lab Invest. 1982 Jul;47(1):87–90. [PubMed] [Google Scholar]
- Haggerty JJ, Jr, Sandler R. Strongyloidiasis presenting as depression: a case report. J Clin Psychiatry. 1982 Aug;43(8):340–341. [PubMed] [Google Scholar]
- Handler AM, Maroy P. Ecdysteroid receptors in Drosophila melanogaster adult females. Mol Cell Endocrinol. 1989 May;63(1-2):103–109. [PubMed] [Google Scholar]
- Harper JS, 3rd, Genta RM, Gam A, London WT, Neva FA. Experimental disseminated strongyloidiasis in Erythrocebus patas. I. Pathology. Am J Trop Med Hyg. 1984 May;33(3):431–443. [PubMed] [Google Scholar]
- Harris RA, Jr, Musher DM, Fainstein V, Young EJ, Clarridge J. Disseminated strongyloidiasis. Diagnosis made by sputum examination. JAMA. 1980 Jul 4;244(1):65–66. [PubMed] [Google Scholar]
- Higenbottam TW, Heard BE. Opportunistic pulmonary strongyloidiasis complicating asthma treated with steroids. Thorax. 1976 Apr;31(2):226–233. [PMC free article] [PubMed] [Google Scholar]
- Igra-Siegman Y, Kapila R, Sen P, Kaminski ZC, Louria DB. Syndrome of hyperinfection with Strongyloides stercoralis. Rev Infect Dis. 1981 May-Jun;3(3):397–407. [PubMed] [Google Scholar]
- Kane MG, Luby JP, Krejs GJ. Intestinal secretion as a cause of hypokalemia and cardiac arrest in a patient with strongyloidiasis. Dig Dis Sci. 1984 Aug;29(8):768–772. [PubMed] [Google Scholar]
- Kennedy S, Campbell RM, Lawrence JE, Nichol GM, Rao DM. A case of severe Strongyloides stercoralis infection with jejunal perforation in an Australian ex-prisoner-of-war. Med J Aust. 1989 Jan 16;150(2):92–93. [PubMed] [Google Scholar]
- Klein RA, Cleri DJ, Doshi V, Brasitus TA. Disseminated Strongyloides stercoralis: a fatal case eluding diagnosis. South Med J. 1983 Nov;76(11):1438–1440. [PubMed] [Google Scholar]
- Kuberski TT, Gabor EP, Boudreaux D. Disseminated strongyloidiasis. A complication of the immunosuppressed host. West J Med. 1975 Jun;122(6):504–508. [PMC free article] [PubMed] [Google Scholar]
- Lambroza A, Dannenberg AJ. Eosinophilic ascites due to hyperinfection with Strongyloides stercoralis. Am J Gastroenterol. 1991 Jan;86(1):89–91. [PubMed] [Google Scholar]
- Lansoud-Soukate J, Gharib B, Baswaid S, Capron A, de Reggi M. Ecdysteroid-like compounds in the serum and urine of African patients infected with Loa loa and Mansonella perstans microfilariae. Trans R Soc Trop Med Hyg. 1990 Mar-Apr;84(2):269–271. [PubMed] [Google Scholar]
- Leelarasamee A, Nimmannit S, Na Nakorn S, Aswapokee N, Aswapokee P, Benjasuratwong Y. Disseminated strongyloidiasis: report of seven cases. Southeast Asian J Trop Med Public Health. 1978 Dec;9(4):539–542. [PubMed] [Google Scholar]
- López JE, Marcano-Torres M, Peña JR, Quintini A, Malpica CC, López JE, López Salazar Y. Hepatitis granulomatosa producida por el Strongyloides stercoralis. Presentación de un caso con confirmación histopatológica. G E N. 1984 Jul-Dec;38(3-4):133–143. [PubMed] [Google Scholar]
- Lucas SB. Missing infections in AIDS. Trans R Soc Trop Med Hyg. 1990;84 (Suppl 1):34–38.[PubMed] [Google Scholar]
- Maayan S, Wormser GP, Widerhorn J, Sy ER, Kim YH, Ernst JA. Strongyloides stercoralis hyperinfection in a patient with the acquired immune deficiency syndrome. Am J Med. 1987 Nov;83(5):945–948. [PubMed] [Google Scholar]
- McLarnon M, Ma P. Brain stem glioma complicated by Strongyloides stercoralis. Ann Clin Lab Sci. 1981 Nov-Dec;11(6):546–549. [PubMed] [Google Scholar]
- McRury J, De Messias IT, Walzer PD, Huitger T, Genta RM. Specific IgE responses in human strongyloidiasis. Clin Exp Immunol. 1986 Sep;65(3):631–638. [PMC free article] [PubMed] [Google Scholar]
- Mercer JG, Munn AE, Arme C, Rees HH. Analysis of ecdysteroids in different developmental stages of Hymenolepis diminuta. Mol Biochem Parasitol. 1987 Aug;25(1):61–71. [PubMed] [Google Scholar]
- Milder JE, Walzer PD, Kilgore G, Rutherford I, Klein M. Clinical features of Strongyloides stercoralis infection in an endemic area of the United States. Gastroenterology. 1981 Jun;80(6):1481–1488. [PubMed] [Google Scholar]
- Milner PF, Irvine RA, Barton CJ, Bras G, Richards R. Intestinal malabsorption in Strongyloides stercoralis infestation. Gut. 1965 Dec;6(6):574–581. [PMC free article] [PubMed] [Google Scholar]
- Morgan JS, Schaffner W, Stone WJ. Opportunistic strongyloidiasis in renal transplant recipients. Transplantation. 1986 Nov;42(5):518–524. [PubMed] [Google Scholar]
- Moura H, Fernandes O, Viola JP, Silva SP, Passos RH, Lima DB. Enteric parasites and HIV infection: occurrence in AIDS patients in Rio de Janeiro, Brazil. Mem Inst Oswaldo Cruz. 1989 Oct-Dec;84(4):527–533. [PubMed] [Google Scholar]
- Murray JF, Garay SM, Hopewell PC, Mills J, Snider GL, Stover DE. NHLBI workshop summary. Pulmonary complications of the acquired immunodeficiency syndrome: an update. Report of the second National Heart, Lung and Blood Institute workshop. Am Rev Respir Dis. 1987 Feb;135(2):504–509. [PubMed] [Google Scholar]
- Nakada K, Yamaguchi K, Furugen S, Nakasone T, Nakasone K, Oshiro Y, Kohakura M, Hinuma Y, Seiki M, Yoshida M, et al. Monoclonal integration of HTLV-I proviral DNA in patients with strongyloidiasis. Int J Cancer. 1987 Aug 15;40(2):145–148. [PubMed] [Google Scholar]
- Nera FA, Murphy EL, Gam A, Hanchard B, Figueroa JP, Blattner WA. Antibodies to Strongyloides stercoralis in healthy Jamaican carriers of HTLV-1. N Engl J Med. 1989 Jan 26;320(4):252–253. [PubMed] [Google Scholar]
- Newton RC, Limpuangthip P, Greenberg S, Gam A, Neva FA. Strongyloides stercoralis hyperinfection in a carrier of HTLV-I virus with evidence of selective immunosuppression. Am J Med. 1992 Feb;92(2):202–208. [PubMed] [Google Scholar]
- Noodleman JS. Eosinophilic appendicitis. Demonstration of Strongyloides stercoralis as a causative agent. Arch Pathol Lab Med. 1981 Mar;105(3):148–149. [PubMed] [Google Scholar]
- Petithory JC, Derouin F. AIDS and strongyloidiasis in Africa. Lancet. 1987 Apr 18;1(8538):921–921. [PubMed] [Google Scholar]
- Pillay SV. Hyperinfection with Strongyloides stercoralis. A report of 3 cases. S Afr Med J. 1978 Oct 14;54(16):670–672. [PubMed] [Google Scholar]
- Poltera AA, Katsimbura N. Granulomatous hepatitis due to Strongyloides stercoralis. J Pathol. 1974 Aug;113(4):241–246. [PubMed] [Google Scholar]
- Purtilo DT, Meyers WM, Connor DH. Fatal strongyloidiasis in immunosuppressed patients. Am J Med. 1974 Apr;56(4):488–493. [PubMed] [Google Scholar]
- Quiñones Soto RA, Harrington PT, Gutiérrez Núez JJ, Ramírez Ronda CH, Bermúdez RH. Estrongiloidiasis en el paciente inmunocomprometido. Bol Asoc Med P R. 1981 Nov;73(11):562–566. [PubMed] [Google Scholar]
- René E, Marche C, Régnier B, Saimot AG, Vittecoq B, Matheron S, Le Port C, Bricaire F, Bure A, Brun-Vezinet C, et al. Manifestations digestives du syndrome d’immunodéficience acquise (SIDA): étude chez 26 patients. Gastroenterol Clin Biol. 1985 Apr;9(4):327–335. [PubMed] [Google Scholar]
- Rivera E, Maldonado N, Vélez-García E, Grillo AJ, Malaret G. Hyperinfection syndrome with Strongyloides stercoralis. Ann Intern Med. 1970 Feb;72(2):199–204. [PubMed] [Google Scholar]
- Schad GA. Cyclosporine may eliminate the threat of overwhelming strongyloidiasis in immunosuppressed patients. J Infect Dis. 1986 Jan;153(1):178–178. [PubMed] [Google Scholar]
- Schad GA, Aikens LM, Smith G. Strongyloides stercoralis: is there a canonical migratory route through the host? J Parasitol. 1989 Oct;75(5):740–749. [PubMed] [Google Scholar]
- Schad GA, Hellman ME, Muncey DW. Strongyloides stercoralis: hyperinfection in immunosuppressed dogs. Exp Parasitol. 1984 Jun;57(3):287–296. [PubMed] [Google Scholar]
- Scowden EB, Schaffner W, Stone WJ. Overwhelming strongyloidiasis: an unappreciated opportunistic infection. Medicine (Baltimore) 1978 Nov;57(6):527–544. [PubMed] [Google Scholar]
- Seth V, Beotra A. Malnutrition and immune system. Indian Pediatr. 1986 Apr;23(4):277–302.[PubMed] [Google Scholar]
- Setia U, Bhatia G. Pancreatic cystadenocarcinoma associated with strongyloides. Am J Med. 1984 Jul;77(1):173–175. [PubMed] [Google Scholar]
- Shelhamer JH, Neva FA, Finn DR. Persistent strongyloidiasis in an immunodeficient patient. Am J Trop Med Hyg. 1982 Jul;31(4):746–751. [PubMed] [Google Scholar]
- da Silva OA, Amaral CF, da Silveira JC, López M, Pittella JE. Hypokalemic respiratory muscle paralysis following Strongyloides stercoralis hyperinfection: a case report. Am J Trop Med Hyg. 1981 Jan;30(1):69–73. [PubMed] [Google Scholar]
- Smith JD, Goette DK, Odom RB. Larva currens. Cutaneous strongyloidiasis. Arch Dermatol. 1976 Aug;112(8):1161–1163. [PubMed] [Google Scholar]
- Vieyra-Herrera G, Becerril-Carmona G, Padua-Gabriel A, Jessurun J, Alonso-de Ruiz P. Strongyloides stercoralis hyperinfection in a patient with the acquired immune deficiency syndrome. Acta Cytol. 1988 Mar-Apr;32(2):277–278. [PubMed] [Google Scholar]
- von Kuster LC, Genta RM. Cutaneous manifestations of strongyloidiasis. Arch Dermatol. 1988 Dec;124(12):1826–1830. [PubMed] [Google Scholar]
- West BC, Wilson JP. Subconjunctival corticosteroid therapy complicated by hyperinfective strongyloidiasis. Am J Ophthalmol. 1980 Jun;89(6):854–857. [PubMed] [Google Scholar]
- Williford ME, Foster WL, Jr, Halvorsen RA, Thompson WM. Emphysematous gastritis secondary to disseminated stronglyloidiasis. Gastrointest Radiol. 1982;7(2):123–126. [PubMed] [Google Scholar]
- Willis AJ, Nwokolo C. Steroid therapy and strongyloidiasis. Lancet. 1966 Jun 25;1(7452):1396–1398. [PubMed] [Google Scholar]
- Yamaguchi K, Matutes E, Catovsky D, Galton DA, Nakada K, Takatsuki K. Strongyloides stercoralis as candidate co-factor for HTLV-I-induced leukaemogenesis. Lancet. 1987 Jul 11;2(8550):94–95. [PubMed] [Google Scholar]
What if M.S, Grand Mal seizures, meningitis and the like are all a result of the central nervous systems protective barrier being destroyed by disseminated Strongyloidiasis? Specifically Larva currens/Cutaneous strongyloidiasis maybe?
From www.epilepsy.com <–THIS – IF YOU HAVE HAD SEIZURES
Strongyloides stercoralis is a small nematode that can parasitize the small bowel of humans. Larvae living freely in moist soil invade rapidly through contacted skin and migrate into lymphatics to reach the venous system, where they travel to the lungs, migrate up airways to the glottis, and then down the esophagus to the small intestine.
When immune function is compromised (e.g., in HIV infection or AIDS), the CNS can become involved in disseminated strongyloidiasis. CNS manifestations can be secondary to larvae infestation. More commonly, however, gut bacteria transmitted by the migrating parasite produce bacterial meningitis. Seizures can be an epiphenomenon of these complications.
Thiabendazole can be helpful if started early in the disease process, but disseminated strongyloidiasis is usually fatal.Comorbid seizure management is routine.162
Adapted from: Goldstein MA and Harden CL. Infectious states. In: Ettinger AB and Devinsky O, eds. Managing epilepsy and co-existing disorders. Boston: Butterworth-Heinemann; 2002;83-133.
With permission from Elsevier (www.elsevier.com).