Acknowledgments
I would like to express my thanks to the many colleagues and students who have helped shape my perspective regarding the field of medical parasitology over the years, especially Rodney D. Adam, Yost Amrein, Bruce Anderson, Michael J. Arrowood, Lawrence Ash, M. Teresa Audicana, Kevin Baird, Gordon Ball, Ralph Barr, J. L. Barratt, Brent Barrett, Marilyn Bartlett, Zvi Bentwich, Henry Bishop, Byron L. Blagburn, Kenneth Borchardt, Peter Boreham, Emilio Bouza, Richard Bradbury, Thomas Brewer, David Bruckner, Sandra Bullock-Iacullo, Andrew Butcher, Guy Cabral, Ann Cali, Vita Cama, Sandy Cairncross, Elizabeth Canning, David Casemore, Francis Chan, Ray Chan, Francois Chappuis, James Cherry, Peter L. Chiodini, John Christie, Frank E. G. Cox, John Cross, Deirdre Church, C. Graham Clark, D. P. Clark, William E. Collins, Peter G. Contacos, Frank Cox, Janet Cox-Singh, S. L. Croft, William Current, A. J. da Silva, Peter Deplazes, Paul DeRycke, Dickson Despommier, L. S. Diamond, J. P. Dubey, Mark Eberhard, Johannes Eckert, David Ellis, J. T. Ellis, Gaétan Faubert, Ronald Fayer, Sydney Finegold, Joe Forsyth, Jacob Frenkel, Thomas Fritsche, H. H. Garcia, Omai Garner, P. J. Gavin, S. Geerts, Robert Goldsmith, Bruno Gottstein, Thaddeus K. Graczyk, Carlos Graeff-Teixeira, Martin P. Grobusch, David A. Haake, Paul Hackendorf, Pauline Hahn, Thomas Hanscheid, Rachidul Haque, J. Harkness, William J. Hausler, George Healy, Barbara Herwaldt, Donald Heyneman, George Hillyer, S. L. Hoffman, Peter Horen, Peter Hotez, Henry D. Isenberg, David John, E. H. Johnson, Stephanie Johnston, Irving Kagan, Ray Kaplan, Kevin R. Kazacos, John Kessel, Mary Klassen-Fischer, Evelyn Kokoskin, Elmer Koneman, W. A. Krotoski, Jaime LaBarca, David A. Leiby, William Lewis, David Lindsay, Andrea Linscott, M. D. Little, Earl Long, Edward Markell, D. Marriott, Marilyn Marshall, A. Mathis, Blaine Mathison, Alan J. Magill, Francine Marciano-Cabral, A. M. Marty, Alexander Mathis, James H. McKerrow, James McLaughlin, Diane McMahon-Pratt, Donald P. McManus, Mae Melvin, Michael Miller, Anthony Moody, Clinton K. Murray, William Murray, Ronald Neafie, Ron Neimeister, Ann Nelson, Susan Novak, Thomas Nutman, Thomas Orihel, Ynes Ortega, Robert Owen, Josephine Palmer, Graeme Paltridge, Liron Pantanowitz, Mark Perry, David Persing, William A. Petri, Jr., Michael Pfaller, N. J. Pieniazek, Dylan R. Pillai, Kathy Powers, E. Pozio, Bobbi Pritt, Paul Prociv, Gary W. Procop, Fred Rachford, Sharon L. Reed, Barth Reller, Julie Ribes, Andrew Rocha, William Rogers, Jon Rosenblatt, Allen G. P. Ross, Norbert Ryan, Judy Sakanari, Carol Santaloci, Peter Schantz, Frederick L. Schuster, James Seidel, Nicholas Serafy, J. A. Shadduck, Harsha Sheorey, Irwin Sherman, Robyn Shimizu, Balbir Singh, James Smith, Rosemary Soave, Frank J. Sorvillo, S. L. Stanley, Jr., John Steele, Deborah Stenzel, Damian Stark, Linda Stetzenbach, Charles Sterling, James J. Sullivan, Alex Sulzer, Kevin S. W. Tan, Egbert Tannich, Herbert Tanowitz, Mehmet Tanyuksel, William Trager, Peter Traynor, Antonio R. L. Teixeira, Sam Telford, William Trager, Allan R. Truant, Jerrold Turner, Saul Tzipori, Jacqueline A. Upcroft, Peter Upcroft, Tom van Gool, Eric Vanderslice, Jacob Verweij, Govinda Visvesvara, Marietta Voge, Susanne Wahlquist, Kenneth Walls, Rainer Weber, Wilfred Weinstein, Louis Weiss, P. P. Wilkins, John Williams, John Wilson, Marianna Wilson, Jeffrey J. Windsor, Washington Winn, Martin Wolfe, Donna Wolk, Johnson Wong, Lihua Xiao, Nigel Yeates, Judy Yost, Wenbao Zhang, Charles and Wiladene Zierdt, and many others whom I may have failed to mention specifically. If the information contained in this edition provides help to those in the field of microbiology, I will have succeeded in passing on this composite knowledge to the next generation of students and teachers.
Special thanks go to Sharon Belkin for her additional illustrations for this edition. I also thank Ronald Neafie from the Armed Forces Institute of Pathology for providing many photographs to illustrate several areas of the book, particularly the information on histological identification of parasites, and Herman Zaiman for providing slides that he has prepared and/or edited from many contributors worldwide. Very special thanks go to the group at the Centers for Disease Control and Prevention for the use of many of their clinical parasitology images; these images are invaluable to the microbiology community and include images contributed to CDC by many others, as well.
I would like to thank members of the editorial staff of ASM Press, especially Ellie Tupper; they are outstanding professionals and made my job not only challenging but fun.
Above all, my very special thanks go to my late husband, John, for his love and support for the many projects that I have been involved in over the years. I could never have undertaken these challenges without his help and understanding, a true partnership.
APPENDIX 6
Reference Sources
GENERAL REFERENCES
1. Ash LR, Orihel TC. 2007. Atlas of Human Parasitology, 5th ed. ASCP Press, Chicago, IL.
2. Ash LR, Orihel TC. 1991. Parasites: A Guide to Laboratory Procedures and Identification. ASCP Press, Chicago, IL.
3. Ash LR, Orihel TC, Savioli L. 1994. Bench Aids for the Diagnosis of Intestinal Parasites. World Health Organization, Geneva, Switzerland.
4. Beaty BJ, Marquardt WC (ed). 1996. The Biology of Disease Vectors. University Press of Colorado, Niwot.
5. Beaver PC, Jung RC, Cupp EW. 1984. Clinical Parasitology, 9th ed. Lea & Febiger, Philadelphia, PA.
6. Binford CH, Connor DH. 1976. Pathology of Tropical and Extraordinary Diseases. Armed Forces Institute of Pathology, Washington, DC.
7. Borchardt KA, Noble MA (ed). 1997. Sexually Transmitted Diseases. CRC Press, Boca Raton, FL.
8. Chernin E. 1977. A bicentennial sampler: milestones in the history of tropical medicine and hygiene. Am J Trop Med Hyg 26:1053–1104.
9. Cook GC, Zumla A (ed). 2008. Manson’s Tropical Diseases, 22nd ed. WB Saunders Co, Philadelphia, PA.
10. Cox FEG, Wakelin D, Gillespie SH, Despommier DD (ed). 2006. Parasitology, vol 5, Topley & Wilson’s Microbiology and Microbial Infections, 10th ed. Edward Arnold, London, United Kingdom.
11. Del Brutto O, Garcia HH. 2014. Cysticercosis of the Human Nervous System. Springer-Verlag, Berlin, Germany.
12. Despommier DD, Gwadz RW, Hotez PJ (ed). 2005. Parasitic Diseases, 5th ed. Springer-Verlag, New York, NY.
13. Eldridge BF, Edman JD (ed). 2004. Medical Entomology, 2nd ed. Kluwer Academic Publishers, Norwell, MA.
14. Farrar J, Hotez PJ, Junghanss T, Kang G, Laloo D, White NJ. 2013. Manson’s Tropical Diseases, 23rd ed. Elsevier Health Sciences, London, United Kingdom.
15. Faust EC. 1949. Human Helminthology, 3rd ed. Lea & Febiger, Philadelphia, PA.
16. Fayer R (ed). 2007. Cryptosporidium and Cryptosporidiosis, 2nd ed. CRC Press, Boca Raton, FL.
17. Foster WD. 1965. A History of Parasitology. E & S Livingstone, London, United Kingdom.
18. Garcia HH, Tanowitz HB, Del Brutto OH. 2013. Neuroparasitology and Tropical Neurology, 3rd ed. Elsevier, Oxford, United Kingdom.
19. Garcia LS. 2009. Practical Guide to Diagnostic Parasitology, 2nd ed. ASM Press, Washington, DC.
20. Goddard J. 2012. Physician’s Guide to Arthropods of Medical Importance, 6th ed. CRC Press, Boca Raton, FL.
21. Goddard J. 2012. Public Health Entomology. CRC Press, Boca Raton, FL.
22. Harwood RF, James MT. 1979. Entomology in Human and Animal Health. Macmillan, New York, NY.
23. Hoeppli R. 1962. Parasites and Parasitic Infections in Early Medicine and Science. University of Malaya Press, Singapore.
24. Horsburgh CR, Jr, Nelson AM (ed). 1997. Pathology of Emerging Infections. ASM Press, Washington, DC.
25. Horsfall WR. 1962. Medical Entomology: Arthropods and Human Disease. The Ronald Press, New York, NY.
26. John DT, Petri WA. 2006. Medical Parasitology, 9th ed. WB Saunders Co, Philadelphia, PA.
27. Jong EC, McMullen R. 2003. The Travel and Tropical Medicine Manual, 3rd ed. W. Saunders Co, Philadelphia, PA.
28. Kean BH, Mott KE, Russell AJ. 1978. Tropical Medicine and Parasitology Classic Investigations. Cornell University Press, Ithaca, NY.
29. Kettle DS. 1995. Medical and Veterinary Entomology, 2nd ed. CAB International, Wallingford, United Kingdom.
30. Klassen-Fischer MK, Meyers WM, Neafie RC. 2011. Topics on the Pathology of Protozoan and Invasive Arthropod Diseases: African Trypanosomiasis. Uniformed Services University of the Health Sciences, Bethesda, MD.
31. Kokoskin E. 2001. The Malaria Manual. McGill University Centre for Tropical Diseases, Montreal, Quebec, Canada.
32. Lambert HP, Farrar WE. 1982. Infectious Diseases Illustrated. WB Saunders Co, Philadelphia, PA.
33. Leventhal R, Cheadle RF. 2012. Medical Parasitology: A Self-Instructional Text, 6th ed. FA Davis, Philadelphia, PA.
34. Lujan HD, Svard SG (ed). 2011. Giardia: A Model Organism. Springer-Verlag, Vienna, Austria.
35. Magill AJ, Meyers WM, Neafie RC, Klassen-Fischer MK. 2011. Topics on the Pathology of Protozoan and Invasive Arthropod Diseases: Cutaneous Leishmaniasis. Uniformed Services University of the Health Sciences, Bethesda, MD.
36. Magill AJ, Meyers WM, Klassen-Fischer MK, Neafie RC. 2011. Topics on the Pathology of Protozoan and Invasive Arthropod Diseases: Visceral Leishmaniasis. Uniformed Services University of the Health Sciences, Bethesda, MD.
37. Magill AJ, Ryan ET, Sullivan T, Hill DR. 2012. Hunter’s Tropical Medicine, 9th ed. WB Saunders Co, Philadelphia, PA.
38. Mansour TE, Mansour JM. 2011. Chemotherapeutic Targets in Parasites. Cambridge University Press, Cambridge, United Kingdom.
39. Mehlhorn H, Tan KSW, Yoshikawa H. 2013. Blastocystis: Pathogen or Passenger? Springer Verlag, Berlin, Germany.
40. Meyers WM, Neafie RC, Marty AM, Wear DJ (ed). 2000. Pathology of Infectious Diseases, vol 1, Helminthiases. Armed Forces Institute of Pathology, Washington, DC.
41. Murray PR, Baron EJ, Jorgensen JH, Landry ML, Pfaller MA (ed). 2007. Manual of Clinical Microbiology, 9th ed. ASM Press, Washington, DC.
42. Murrell KD, Fried B (ed). 2007. Food-Borne Parasitic Zoonoses. Springer-Verlag, New York, NY.
43. Nelson AM, Horsburgh DR Jr. 1998. Pathology of Emerging Infections 2. ASM Press, Washington, DC.
44. Orihel TC, Ash LR. 1995. Parasites in Human Tissues. ASCP Press, Chicago, IL.
45. Orihel TC, Ash LR, Ramachandran CP. 1996. Bench Aids for the Diagnosis of Filarial Infections. World Health Organization, Geneva, Switzerland.
46. Ortega YR. 2006. Foodborne Parasites. Springer-Verlag, New York, NY.
47. Peters W, Gilles HM. 1995. Color Atlas of Tropical Medicine and Parasitology. Mosby-Wolfe, New York, NY.
48. Reeder MM, Palmer PES. 1981. The Radiology of Tropical Diseases with Epidemiological, Pathological and Clinical Correlation. Williams & Wilkins Co, Baltimore, MD.
49. Roberts LS, Janovy J, Nadler S. 2012. Foundations of Parasitology, 9th ed. CV Mosby Co, St. Louis, MO.
50. Rondanelli EG, Scaglia M. 1993. Atlas of Human Protozoa. Masson, Milan, Italy.
51. Scarpignato C, Rampal P (ed). 1995. Travelers’ Diarrhea: Recent Advances. Karger Publishers, Farmington, CT.
52. Scott HH. 1939. A History of Tropical Medicine. Edward Arnold & Co, London, United Kingdom.
53. Secor WE, Colley DG (ed). 2010. Schistosomiasis. Springer-Verlag, New York, NY.
54. Sherman IW (ed). 1998. Malaria. ASM Press, Washington, DC.
55. Sherman IW. 2008. Reflections on a Century of Malaria Biochemistry. Academic Press, Inc. San Diego, CA.
56. Smith KCV. 1973. Insects and Other Arthropods of Medical Importance. John Wiley & Sons, Inc, New York, NY.
57. Spencer FM, Monroe LS. 1976. The Color Atlas of Intestinal Parasites, 2nd ed. Charles C Thomas, Publisher, Springfield, IL.
58. Staines HM, Krishna S (ed). 2012. Treatment and Prevention of Malaria. Birkhauser Verlag AG, Basel, Switzerland.
59. Steele JH. 1982. CRC Handbook Series in Zoonoses. CRC Press, Inc, Boca Raton, FL.
60. Sullivan D, Krishna S (ed). 2010. Malaria: Drugs, Disease and Post-Genomic Biology. Springer-Verlag, Berlin, Germany.
61. Sun T. 1999. Parasitic Disorders, 2nd ed. Williams & Wilkins Co., Baltimore, MD.
62. Taylor AER, Baker JR. 1978. Methods of Cultivating Parasites In Vitro. Academic Press Inc., New York, NY.
63. Taylor MA, Coop RL, Wall RL. 2007. Veterinary Parasitology, 3rd ed. Blackwell Publishing, Chichester, United Kingdom.
64. Von Brand T. 1979. Biochemistry and Physiology of Endoparasites. Elsevier/North Holland Publishing Co, New York, NY.
65. Warren KS, Mahmoud AAF. 1990. Tropical and Geographical Medicine, 2nd ed. McGraw Hill Book Co, New York, NY.
66. Wittner M (ed), Weiss LM (contributing ed). 1999. The Microsporidia and Microsporidiosis. ASM Press, Washington, DC.
67. Yamaguchi T. 1981. Color Atlas of Clinical Parasites. Lea & Febiger, Philadelphia, PA.
68. Zaman V. 1994. Atlas of Medical Parasitology, 3rd ed. Lea & Febiger, Philadelphia, PA.
JOURNALS AND PERIODICALS
Acarologia, 1959 to date.
Acta Tropica, 1944 to date.
Advances in Parasitology, 1963 to date.
American Journal of Tropical Medicine, 1921 to 1951.
American Journal of Tropical Medicine and Hygiene, 1952 to date.
Annals of the Entomological Society of America, 1908 to date.
Pathogens and Global Health (Annals of Tropical Medicine and Parasitology), 1907 to date.
Annual Review of Entomology, 1956 to date.
Bulletin of Entomological Research, 1910 to date.
Clinical Microbiology Reviews, 1988 to date.
Emerging Infectious Diseases, 1995 to date.
Experimental Parasitology, 1951 to date.
International Journal for Parasitology, 1971 to date.
Journal of Clinical Microbiology, 1963 to date.
Journal of Helminthology, 1923 to date.
Journal of Medical Entomology, 1964 to date.
Journal of Tropical Medicine and Hygiene, 1898 to date.
Malaria Journal, 2009 to date.
Memórias do Instituto Oswaldo Cruz, 1909 to date.
Molecular and Biochemical Parasitology, 1980 to date.
Mosquito News, 1941 to date.
Mosquito Systematics, 1969 to date.
Parasite Immunology, 1979 to date.
Parasitology, 1908 to date.
Parasitology Research, 1987 to date.
Parasitology Today, 1985 to date.
PLoS Neglected Tropical Diseases, 2008 to date.
Proceedings of the Helminthological Society of Washington, 1934 to date.
Puerto Rico Journal of Public Health and Tropical Medicine, 1925 to 1950.
Systematic Parasitology, 1979 to date.
The Journal of Eukaryotic Microbiology, 1993 to date.
Transactions of the American Microscopical Society, 1892 to date.
Transactions of the Royal Society of Tropical Medicine and Hygiene, 1907 to date.
Trends in Parasitology, 2001 to date.
Tropical and Geographical Medicine, 1949 to date (continuation of Documenta de Medicina Geographica et Tropica).
Veterinary Parasitology, 1975 to date.
World Health Organization Monographs, Technical Reports, Reports of Expert Committees, Bulletin, and Chronicle, 1948 to date.
ABSTRACTS AND BIBLIOGRAPHIC SOURCES
Biological Abstracts, 1929 to date.
Excerpta Medica, section 4, Microbiology, 1948 to date.
Helminthological Abstracts, 1932 to date.
Index Catalogue of Medical and Veterinary Zoology (author catalog), 1932 to 1952 with current supplements.
Index Medicus, 1878 to 1889, 1903 to 1927, 1960 to date.
Protozoological Abstracts, 1977 to date.
Quarterly Cumulative Index, 1916 to 1926, 1927 to 1956.
Quarterly Bibliography of Major Tropical Diseases, 1978 to date.
Review of Applied Entomology, section B (Medical and Veterinary), 1913 to date.
Tropical Diseases Bulletin, 1913 to date.
WEBSITES
Parasite Images
Do a web search under general parasite categories and/or specific genus/species of parasites, then select “images”; many images will appear. However, you may see the same image under different sites/links; therefore, it is difficult to determine which site is the original. This impacts your ability to use the images due to possible copyright issues.
Parasitology Information
Centers for Disease Control and Prevention:
www.dpd.cdc.gov/dpx/
Medical Chemical Corporation:
www.med-chem.com
Literature
NCBI National Library of Medicine (PubMed):
http://www.ncbi.nlm.nih.gov/pubmed
Government Websites Related to Health Care Regulations
Centers for Medicare and Medicaid Services (CMS):
http://cms.hhs.gov/
CMS Fee Schedules:
http://www.cms.gov/apps/physician-fee-schedule/license-agreement.aspx
CMS Program Integrity Issues:
http://www.cms.gov/Medicare-Medicaid-Coordination/Fraud-Prevention/MedicaidIntegrityProgram/index.html?redirect=/medicaidintegrityprogram/
Compliance Program, Fraud Alerts, Advisory Opinions, Red Book, Work Plan:
http://oig.hhs.gov/
Medicare Learning Network:
http://www.cms.gov/Outreach-and-Education/Medicare-Learning-Network-MLN/MLNGenInfo/
National Technical Information Service:
www.ntis.gov
National Center for Health Statistics:
http://www.cdc.gov/nchs/about.htm
Office of the Inspector General (OIG) Compliance Documents for Clinical Laboratories, Hospitals, and Third Party Billing:
http://www.gpo.gov/fdsys/search/search.action?st=Compliance+document+for+clinical+laboratories%2C+hospitals+and+%22third+party%22+billing
APPENDIX 7
“Late-Breaking” Published Information
The following information was published in the last couple of months of 2014 and the first few months of 2015 and contains relevant information for the field of diagnostic medical parasitology. This is “late-breaking” published information that may be helpful for the reader.
Intestinal Protozoa (Amebae)
Entamoeba spp.
Petropolis DB, Faust DM, Deep Jhingan G, Guillen N. 2014. A new human 3D-liver model unravels the role of galectins in liver infection by the parasite Entamoeba histolytica. PLoS Pathog 10:e1004381. doi:10.1371/journal.ppat.1004381. PMID 25211477
The authors developed a human three-dimensional (3D) liver in vitro model using cultured liver sinusoidal endothelial cells and hepatocytes in a collagen-I matrix. They found important hepatic markers and showed that the cell layers act as a biological barrier. Invasion of Entamoeba histolytica was investigated using wild-type strains and amebae with different virulence or adhesive properties. They demonstrated that amebic Gal/GalNAc lectin influenced the dependence of the endothelium crossing. This 3D liver model supported the molecular analysis of human cellular responses and showed for the first time the importance of the role of human galectins in the adhesion of parasites to the endothelial cells. The presence of galectin-1 and -3 in the extracellular medium led to proinflammatory cytokine release, thus supporting the role for human galectins’ hepatic inflammatory response. The approach in this study using a 3D liver model may be relevant for other parasitic or viral hepatic infections.
Blastocystis spp.
Bálint A, Dóczi I, Bereczki L, Gyulai R, Szücs M, Farkas K, Urbán E, Nagy F, Szepes Z, Wittmann T, Molnár T. 2014. Do not forget the stool examination!: cutaneous and gastrointestinal manifestations of Blastocystis sp. infection. Parasitol Res 113:1585–1590. PMID 24553977
Blastocystis spp. are the most common parasite in the human intestinal tract worldwide. The infection is characterized by diarrhea and abdominal pain; however, extraintestinal findings such as skin lesions may also be present. This study was designed to assess the frequency, clinical symptoms, and skin manifestations of confirmed positive Blastocystis spp. infections. Eighty patients were assessed retrospectively. The average age of the patients was 46.3 years. The number of female patients was higher than the number of males (48 versus 32; 60 versus 40%). Gastrointestinal and dermatological symptoms, biochemical tests, and hematological tests were included in the study. The skin manifestations were analyzed using descriptions, photos, and biopsies. Patients (11.25%) exhibited skin manifestations associated with Blastocystis spp., mainly seen in females. Blastocystis spp. were found in 6% of symptomatic patients between 2005 and 2013. Of the 80 patients with confirmed infection, 73.75% had gastrointestinal symptoms: 40 patients had abdominal pain and 17 had blood in their stool; other symptoms, such as excess gas (15), weight loss (8), perianal pain or itching (6), passing stool with mucus (5), vomiting (2), and fever (2), were less common. Abdominal pain in the group without skin lesions was higher than in patients with skin problems (P = 0.007). The mean value of C-reactive protein demonstrated elevated levels, but eosinophils were normal. Also, there were no significant differences in eosinophilia between patients with versus without skin manifestations. Based on these results, the authors suggest a stool parasite examination for patients with skin lesions of unknown origin.
Ruaux CG, Stang BV. 2014. Prevalence of Blastocystis in shelter-resident and client-owned companion animals in the US Pacific Northwest. PLoS One 9:e107496. doi:10.1371/journal.pone.0107496. PMID 25226285
It has been demonstrated that domestic dogs and cats can be infected with a number of protozoan enteric parasites, including Blastocystis spp. For this reason, there is increased interest in Blastocystis as a potential enteric pathogen and in any possible role dogs and cats may play in human infections. Fecal specimens were collected from 103 shelter-resident dogs, 105 shelter-resident cats, 51 client-owned dogs, and 52 client-owned cats. Blastocystis was detected and subtypes identified using a nested PCR. Shelter animals were much more likely to test positive for Blastocystis (P < 0.05 for dogs, P = 0.009 for cats). Sequence analysis of these animals confirmed that they were carrying a number of Blastocystis subtypes. There was no relationship between Blastocystis infection and gastrointestinal symptoms in either dogs or cats. These data suggest that shelter-companion dogs and cats are more likely to be positive for Blastocystis spp. However, the data also suggest that Blastocystis spp. are unlikely to be major enteric pathogens in these species.
Fréalle E, El Safadi D, Cian A, Aubry E, Certad G, Osman M, Wacrenier A, Dutoit E, Creusy C, Dubos F, Viscogliosi E. 2015. Acute blastocystis-associated appendicular peritonitis in a child, Casablanca, Morocco. Emerg Inf Dis 21:91–94. PMID 25528951
This represents another case in which a Blastocystis sp. has been implicated as a disseminated infection (lumen, mucosa) associated with acute inflammation of the appendix. Although 26 other family members reported cases of acute to moderate gastroenteritis, diagnostic parasitology testing was not performed; thus, the definitive diagnosis in these individuals was not confirmed.
Intestinal Protozoa (Flagellates and Ciliates)
Dientamoeba fragilis
Stark D, Garcia LS, Barratt JL, Phillips O, Roberts T, Marriott D, Harkness J, Ellis JT. 2014. Description of Dientamoeba fragilis cyst and precystic forms from human samples. J Clin Microbiol 52:2680–2683. PMID 24808242
Dientamoeba fragilis is a common pathogenic protozoan of humans and has been reported worldwide. Although the cyst stage of the parasite has been described in an animal model, no cyst stage in humans had been described in detail from clinical samples. In this study, the authors describe the precyst and cyst forms from human clinical samples. In light of the fact that D. fragilis infection can cause chronic long-term infections, these cyst forms may be involved in autoinfection. The nuclear structure is identical to that found in D. fragilis trophozoites. All cysts identified were binucleate, with each nucleus containing the large central karyosome and a delicate nuclear membrane. The nucleus is often fragmented into separate chromatin granules, and chromatin is usually not seen on the nuclear membrane. However, these “true” cysts are rarely seen and identified in clinical samples. The precystic forms were more commonly seen, with a prevalence of up to 5% in human clinical specimens. This precystic form has a compact spherical shape and is up to 50% smaller than “normal” trophozoites. The precysts range in size from 4 to 5 µm, and the cytoplasm is denser and more darkly staining than is seen in normal trophozoites. Also, the cytoplasm rarely contains any inclusions.
Giardia lamblia
Liao JY, Guo YH, Zheng LL, Li Y, Xu WL, Zhang YC, Zhou H, Lun ZR, Ayala FJ, Qu LH. 2014. Both endo-siRNAs and tRNA-derived small RNAs are involved in the differentiation of primitive eukaryote Giardia lamblia. Proc Natl Acad Sci USA 111:14159–14164. PMID 25225396
In eukaryotes, small RNAs (sRNAs) including microRNAs and endogenous siRNAs (endo-siRNAs) regulate most biologic processes such as cell division and differentiation. The authors deep-sequenced the sRNA transcriptome of four different stages in the differentiation of Giardia lamblia, a very primitive eukaryote. They identified a large number of endo-siRNAs and found that they were produced from live telomeric retrotransposons and three genomic regions (i.e., endo-siRNA-generating regions [eSGRs]). Gradual upregulation of endo-siRNAs in the differentiation of Giardia suggests involvement in this process regulation. The authors identified five new kinds of tRNA-derived sRNAs. They also found that the biogenesis in four might correlate with previously discovered stress-induced tRNA-derived RNA (sitRNA). These studies demonstrate a complex group of sRNAs in G. lamblia and further clarify the origin and evolution of eukaryotic sRNAs.
Intestinal Protozoa (Coccidia and Microsporidia)
Cryptosporidium spp.
Yu Y, Zhang H, Guo F, Sun M, Zhu G. 2014. A unique hexokinase in Cryptosporidium parvum, an apicomplexan pathogen lacking the Krebs cycle and oxidative phosphorylation. Protist 165:701–714. PMID 25216472
It is well known that Cryptosporidium parvum may cause untreatable infections in patients with AIDS. This parasite has also been identified as one of the top four diarrheal pediatric pathogens in developing countries. Cryptosporidium is different from other apicomplexans such as Plasmodium and Toxoplasma, since it lacks many metabolic pathways including the Krebs cycle and cytochrome-based respiratory chain and depends on glycolysis for the production of ATP. This study reports on molecular and biochemical characteristics of a hexokinase in C. parvum (CpHK). Phylogeny suggests that apicomplexan hexokinases are highly divergent from those of humans and animals. CpHK differs from those in mammals and Toxoplasma gondii (TgHK) by preferring various hexoses and the capacity to use ATP and other NTPs. While the exact action of 2-deoxy-D-glucose on Cryptosporidium requires confirmation, these data suggest that CpHK and the glycolytic pathway provide potential options for the development of anticryptosporidial therapeutics.
Chandra V, Torres M, Ortega YR. 2014. Efficacy of wash solutions in recovering Cyclospora cayetanensis, Cryptosporidium parvum, and Toxoplasma gondii from basil. J Food Prot 77:1348–1354. PMID 25198596
It has been well documented that parasitic infections can originate from the ingestion of contaminated raw or processed fresh produce (herbs and fruits). Method sensitivity for organism detection on fresh produce is partially dependent on the effectiveness of the wash solutions used. In this study, six wash solutions (sterile E-Pure water, 3% levulinic acid–3% sodium dodecyl sulfate, 1 M glycine, 0.1 M phosphate-buffered saline, 0.1% Alconox, and 1% HCl-pepsin) were evaluated for their ability to remove Cyclospora cayetanensis, Cryptosporidium parvum, and Toxoplasma gondii from basil. Oocysts (100 or 1,000) were inoculated onto the upper surfaces of 25 g of basil leaves, placed in stomacher bags, and stored for 1 h at 21°C or 24 h at 4°C. Leaves were then hand washed in each solution for 1 min. DNA from the various wash solutions was extracted and amplified using PCR for parasite detection. Oocysts (1,000 oocysts per 25 g of basil) were detected in all wash solutions. At a concentration of 100 oocysts per 25 g, oocysts were found in 18.5 to 92.6% of the solutions. The lowest variability in oocyst recovery inoculated with 100 oocysts was seen in the 1% HCl-pepsin wash solution. Oocyst recovery was greater at 1 h than at 24 h postinoculation. This study provides guidance for the use of wash solutions for produce, providing the most efficient retrieval of parasite oocysts.
Ryan U, Fayer R, Xiao L. 2014. Cryptosporidium species in humans and animals: current understanding and research needs. Parasitology 141:1667–1685. PMID 25111501
Cryptosporidium is widely recognized as a major cause of diarrhea in developing countries. Since treatment options are very limited, control depends on an awareness of the organism biology and transmission. Currently, 26 species are recognized on the basis of morphological, biological, and molecular studies. Of those reported in humans, Cryptosporidium hominis and Cryptosporidium parvum are responsible for the most infections. Livestock, particularly cattle, serve as an important reservoir of zoonotic infections. However, research limitations include the lack of subtyping information for many veterinary Cryptosporidium species. There is limited understanding regarding host specificity of Cryptosporidium. The impact of climate change is also not well defined regarding organism transmission.
Sponseller JK, Griffiths JK, Tzipori S. 2014. The evolution of respiratory cryptosporidiosis: evidence for transmission by inhalation. Clin Microbiol Rev 27:575–586. PMID 24982322
Cryptosporidium infects all major vertebrates and causes significant diarrhea in humans. Children and immunodeficient individuals are more at risk for cryptosporidiosis, especially in developing countries, where high morbidity and mortality figures are seen in preschool-age children. Unfortunately, no antiprotozoal agent or vaccine exists for treatment or prevention. Respiratory cryptosporidiosis has been described in birds and mammals, including immunocompromised humans. Respiratory cryptosporidiosis may also be seen in immunocompetent children with cryptosporidial diarrhea and unexplained cough. In addition to the traditional fecal-oral route of infection, data from animal studies, human clinical case reports, and epidemiological studies suggest respiratory transmission via secretions. Transmission of Cryptosporidium oocysts may occur through inhalation or by contact with fomites contaminated by coughing, particularly since the oocysts have been found to survive for some time on fomites. Clarifying the potential role of the respiratory tract in the transmission of Cryptosporidium may help expand the current guidelines for disease prevention.
Chappell CL, Okhuysen PC, Langer-Curry RC, Lupo PJ, Widmer G, Tzipori S. 2015. Cryptosporidium muris: infectivity and illness in healthy adult volunteers. Am J Trop Med Hyg 92:50–55. PMID 25311695
Although human infections with Cryptosporidium muris have been reported in the literature, no definitive information has been available on clinical outcomes in healthy individuals. In this study, six healthy adult volunteers were challenged with 105 viable oocysts and monitored for clinical symptoms. All volunteers became infected, two suffered diarrhea, and three shed oocysts for 7 months (a longer time frame than reported with other species and volunteer infections). Thus, it is clear that healthy individuals are susceptible to infection with C. muris, which causes symptoms and can become a chronic infection for many months.
Microsporidia
Santiana M, Pau C, Takvorian PM, Cali A. 2014. Analysis of the beta-tubulin gene and morphological changes of the microsporidium Anncaliia algerae both suggest albendazole sensitivity. J Eukaryot Microbiol 62:60–68. PMID 25105446
Anncaliia algerae, an obligate intracellular microsporidian parasite, is an opportunistic human pathogen; however, therapy has not been assessed. Albendazole is an antitubulin polymerization drug used for helminth infections and is the drug of choice for some human microsporidial infections. Results vary from organism clearance (Encephalitozoon) to resistance (Enterocytozoon). This study investigated albendazole efficacy on A. algerae infection in rabbit kidney (RK13) cells and human fetal lung (HFL-1) fibroblasts. This drug is effective on A. algerae, and the 50% inhibitory concentration in RK13 cells is 0.1 µg/ml. Long-term therapy can inhibit 98% spore production; however, if the treatment is interrupted, the infection is reestablished without documented additional patient exposures. The A. algerae beta-tubulin gene was purified, cloned, and then sequenced. Specific amino acids linked to benzimidazole sensitivity (five out of six) are conserved in A. algerae. Although these data suggest that A. algerae is sensitive to albendazole, organisms in culture are not completely eliminated.
Hocevar SN, Paddock CD, Spak CW, Rosenblatt R, Diaz-Luna H, Castillo I, Luna S, Friedman GC, Antony S, Stoddard RA, Tiller RV, Peterson T, Blau DM, Sriram RR, da Silva A, de Almeida M, Benedict T, Goldsmith CS, Zaki SR, Visvesvara GS, Kuehnert MJ, Microsporidia Transplant Transmission Investigation Team. 2014. Microsporidiosis acquired through solid organ transplantation: a public health investigation. Ann Intern Med 160:213–220. PMID 24727839
Encephalitozoon cuniculi has been associated with renal, respiratory, and central nervous system infections in immunosuppressed patients. This microsporidian has also been identified as causing illness among three recipients of solid organ transplants from a single donor. Appropriate specimens were examined via culture, immunofluorescent antibody, PCR, immunohistochemistry, and electron microscopy. Medical records from the donor were reviewed, as were answers from a clinically relevant questionnaire. Kidneys and lungs from the donor were transplanted to three recipients. Subsequently, the recipients became symptomatic with fever approximately 7 to 10 weeks posttransplantation. Urine cultures, serologic results, and PCR testing from the three recipients were all positive for E. cuniculi, genotype III. E. cuniculi was also confirmed in all biopsy and autopsy specimens. Although autopsy specimens were not available, serologic test results for E. cuniculi were also found from the donor. The recipients were treated with albendazole. Assessment of the donor information failed to confirm specific factors related to the suspected E. cuniculi infection. Microsporidiosis has now been confirmed as an emerging transplant-associated disease and should be considered in symptomatic transplant recipients. Possible donor-derived disease assessment is critical, especially when multiple recipients from a common donor become symptomatic.
Watts MR, Chan RC, Cheong EY, Brammah S, Clezy KR, Tong C, Marriott D, Webb CE, Chacko B, Tobias V, Outhred AC, Field AS, Prowse MV, Bertouch JV, Stark D, Reddel SW. 2014. Anncaliia algerae microsporidial myositis. Emerg Infect Dis 20:185–191. PMID 24447398
The microsporidian Anncaliia algerae was documented in 2004 as causing fatal myositis in an immunosuppressed patient from Pennsylvania. Two cases were later reported, and the authors reveal two additional cases, including one nonfatal case. All five cases were investigated, including clinical findings, diagnosis, and management; life cycle and epidemiology information was also summarized. Prior to infection, all patients were immunosuppressed due to relevant therapy for rheumatoid arthritis or solid organ transplantation. Four of the five patients were from Australia. The diagnosis was confirmed from skeletal muscle biopsies. The surviving patient was treated with albendazole and had immunosuppressive drugs reduced. While insects serve as the normal hosts for A. algerae, it has been suggested that human contact with water contaminated by A. algerae spores may lead to infections. Therefore, A. algerae has been documented as causing human myositis, particularly in coastal areas in Australia.
Cystoisospora (Isospora)
Lindsay DS, Houk AE, Mitchel SM, Dubey JP. 2014. Developmental biology of Cystoisospora (Apicomplexa: Sarcocystidae) monozoic tissue cysts. J Parasitol 100:392–398. PMID 24841928
Tissue cyst stages form an integral part of the developmental cycle and potential transmission of Sarcocystidae species. Tissue cysts within these species (Toxoplasma, Hammondia, Neospora, Besnoitia, and Sarcocysti) contain many infectious stages (bradyzoites). The tissue cyst stage of Cystoisospora (syn. Isospora) has one infectious stage (zoite) and is therefore referred to as a monozoic tissue cyst (MZTC). No tissue cyst stages are known for the Nephroisospora. The current report looks at the developmental biology of MZTC stages of Cystoisospora Frenkel, 1977, the cause of intestinal coccidiosis in cats, dogs, pigs, and humans. These MZTC stages of Cystoisospora belli are possibly associated with human clinical disease reoccurrence. Developmental stages that are found in low numbers in immunocompetent definitive human hosts tend to be much more numerous in immunosuppressed humans. It is anticipated that additional studies will lead to a clarification of C. belli infections in immunocompromised patients, as well as any life cycle mechanisms associated with possible reoccurrence of disease.
Sarcocystis spp.
Lingappa HA, Krishnamurthy A, Puttaveerachary AK, Govindashetty AM, Sahni S. 2015. Foray of cytologically diagnosed intramuscular sarcocystosis—a rarity. J Clin Diagn Res 9:ED11–ED12. PMID 26155487
While human cases with Sarcocystis spp. are uncommon, these organisms are well documented to cause intestinal and muscular problems. In this particular patient, an intramuscular swelling in the lumbar region was diagnosed through cytology as intramuscular sarcocystosis; this finding was later confirmed from histopathology. This case emphasizes the importance of fine needle aspiration cytology in differentiating sarcocystosis from other intramuscular parasites. In this case, the overall parasite morphology was more definitive from the cytology smears than the traditional paraffin sections and strongly suggests the benefits of using fine needle aspirate cytology. There are excellent images in this particular article.
Fayer R, Esposito DH, Dubey JP. 2015. Human infections with Sarcocystis species. Clin Microbiol Rev 28:295–311. PMID 25715644
Recurrent outbreaks of human cases of muscular sarcocystosis in travelers to Malaysia have led to increased interest in this disease. The Sarcocystis life cycle requires two hosts, definitive and intermediate; humans can serve as definitive hosts with intestinal sarcocystosis (Sarcocystis hominis from beef, and Sarcocystis suihominis from pork). Humans serve as intermediate hosts for Sarcocystis nesbitti (reptile definitive host, and possibly other unidentified species) acquired by ingesting sporocysts from feces-contaminated food or water and the environment. It is important to consider this infection in patients with a relevant travel history to tropical areas of the world, particular if they present with elevated serum enzyme levels and eosinophilia.
Protozoa from Other Body Sites
Free-Living Amebae
Robaei D, Carnt N, Minassian DC, Dart JK. 2014. The impact of topical corticosteroid use before diagnosis on the outcome of Acanthamoeba keratitis. Ophthalmology 121:1383–1388. PMID 24630688
Corticosteroid use prior to the diagnosis of Acanthamoeba keratitis (AK) is strongly linked to a worse visual outcome. This situation often occurs due to the initial misdiagnosis of AK as herpetic keratitis. It is clinically critical to include AK in the differential diagnosis of keratitis causative agents, particularly before diagnosing herpes keratitis and instituting the subsequent use of topical corticosteroids.
Van der Beek NA, van Tienen C, de Haan JE, Roelfsema J, Wismans PJ, van Genderen PJJ, Tanghe HL, Verdijk RM, Titulaer MJ, van Hellemond JJ. 2015. Fatal Balamuthia mandrillaris meningoencephalitis in the Netherlands after travel to The Gambia. Emerg Infect Dis 21:896–898. PMID 25897644
Up to this point, meningoencephalitis caused by Balamuthia mandrillaris had not been reported from Africa. Unlike Acanthamoeba encephalitis, Balamuthia amebic encephalitis has been found in immunocompetent individuals. These findings confirm the virulence of this pathogen. In immunocompetent hosts, an inflammatory response is mounted and amebae are surrounded by macrophages, lymphocytes, and neutrophils. However, with rare exceptions, these patients also tend to die of severe central nervous system disease. Findings are often nonspecific, including an elevated cerebrospinal fluid protein and low cerebrospinal fluid glucose. Infection with this organism should be considered in unexplained cases of meningoencephalitis, including those in immunocompetent patients. In areas of the world where reports are lacking or are minimal, the infection may go unrecognized.
Trichomonas vaginalis
Hathorn E, Ng A, Page M, Hodson J, Gaydos C, Ross JD. 2014. A service evaluation of the Gen-Probe APTIMA nucleic acid amplification test for Trichomonas vaginalis: should it change whom we screen for infection? Sex Transm Infect 91:81–86. PMID 25170162
This study involves evaluation of the Gen-Probe APTIMA nucleic acid amplification test used to determine the prevalence of Trichomonas vaginalis infection in a United Kingdom sexual health clinic. Also, a part of the study was designed to identify possible risk factors to develop an appropriate patient screening strategy. Patients (new clinical episode) were offered the Aptima test as well as routine screening. Asymptomatic females submitted a self-collected vulvovaginal specimen, and asymptomatic men submitted a first-void urine sample. A urethral swab was taken from symptomatic men, and two posterior vaginal swabs from symptomatic females (Aptima and culture). Data were collected from all patients who were positive for T. vaginalis infection, as well as 100 randomly selected T. vaginalis–negative controls. The prevalence of T. vaginalis infection was 21/1,483 (1.4%) (95% confidence interval [CI], 0.9% to 2.2%) in men and 72/2,020 (3.6%) (95% CI, 2.8% to 4.5%) in women. The T. vaginalis positivity rate was higher in black Caribbean patients when compared with Caucasian patients (men 5.4% versus 0.1%, P < 0.001; women 9.0% versus 1.2%, P < 0.001). Compared to culture results, the Aptima test detected 16 more infections (38%) in symptomatic women. Although more inclusive screening with the Aptima test will identify additional T. vaginalis infections, prevalence in the UK is low, and the extra screening may not prove to be cost-effective. However, testing of symptomatic patients and those within high-risk asymptomatic groups may be clinically warranted.
Nathan B, Appiah J, Saunders P, Heron D, Nichols T, Brum R, Alexander S, Baraitser P, Ison C. 2014. Microscopy out performed in a comparison of five methods for detecting Trichomonas vaginalis in symptomatic women. Int J STD AIDS 26:251–256. PMID 24855131
This study was designed to compare five methods (an in-house PCR, the Aptima T. vaginalis kit, the OSOM Trichomonas Rapid Test, culture, and microscopy) to detect Trichomonas vaginalis infection in symptomatic women. A true positive was defined as any sample that was positive for T. vaginalis by two or more methods. Of 246 women, 24 were found positive for T. vaginalis by two or more different methods. From this group of 24 patients, 21 were real-time PCR positive (sensitivity 88%), 22 were Aptima T. vaginalis positive (sensitivity 92%), 22 were OSOM positive (sensitivity 92%), 9 were wet mount microscopy positive (sensitivity 38%), and 21 were culture positive (sensitivity 88%). Two patients were not considered true positives, because they were positive using one method only. All methods demonstrated a sensitivity that was significantly greater than that obtained using wet mount microscopy. Thus, many symptomatic cases are routinely missed if microscopy is the only method used for the diagnosis of trichomoniasis.
Secor WE, Meites E, Starr MC, Workowski KA. 2014. Neglected parasitic infections in the United States: trichomoniasis. Am J Trop Med Hyg 90:800–804. PMID 24808247
Trichomonas vaginalis is the most prevalent nonviral sexually transmitted infection within the United States. However, the problems associated with this infection are not thoroughly understood, and the infection receives too little attention regarding public health issues. Serious infection sequelae include an increased risk of infection with HIV and unfavorable pregnancy outcomes such as premature rupture of the membranes, preterm labor, and low-birth-weight infants. Newly developed diagnostic tests have improved the diagnosis, but infections in males require additional study and clarification. Other issues that need additional investigation include the epidemiology of trichomoniasis in both sexes, defining the actual public health impact of symptomatic and asymptomatic T. vaginalis infections, and therapeutics and their efficacy in reducing the overall costs of this disease. Other important issues for clarification include resistance versus noncompliance with therapy and the probability of reinfection.
Tissue Protozoa
Toxoplasma gondii
McAuley JB. 2014. Congenital toxoplasmosis. J Pediatric Infect Dis Soc 3:S30–S35. PMID 25232475
Infection with Toxoplasma gondii is a very common human parasitic infection; however, patients generally remain asymptomatic. Serious sequelae in a pregnant woman with a primary infection, particularly early in the pregnancy, can cause very serious disease in the developing fetus. These potential problems, including organism potential infectivity and severity of symptoms, have recently been linked to various Toxoplasma genotypes. Risk factors, including foodborne transmission, have been more thoroughly investigated and defined. Early diagnosis in infected pregnant women, followed by recommended therapy, has decreased mother-to-child transmission. The addition of immunoglobulin G avidity testing has also produced improved diagnostic testing and better clinical outcomes during pregnancy. It is now well documented that the infected child can be treated in utero and through the first year of life. However, regardless of our more complete comprehension of congenital disease, other issues remain problematic and require clarification. These include the development of enhanced diagnostic tests, the need for more effective therapy, clearly defined screening protocols for pregnant women, and food safety recommendations for disease prevention.
Davis SM, Anderson BL, Schulkin J, Jones K, Eng JV, Jones JL. 2014. Survey of obstetrician-gynecologists in the United States about toxoplasmosis: 2012 update. Arch Gynecol Obstet 291:545–555. PMID 25205181
The American College of Obstetricians and Gynecologists (ACOG) surveyed a number of their members to establish current knowledge, practices, opinions, and educational preferences regarding toxoplasmosis during pregnancy. Among the respondents, 80.2% had reported no acute maternal Toxoplasma gondii infections during the past 5 years, 12.7% correctly identified the clinical relevance of the Toxoplasma avidity test for screening, and 42.6% performed T. gondii serologies for some asymptomatic pregnant women. Respondents in the northeastern United States were twice as likely to use routine screening when compared with those in the west; female physicians were one and a half times more likely than male physicians to use routine screening. Most practitioners felt that updated ACOG guidelines on screening and management for pregnant women with acute toxoplasmosis would support educational efforts related to this infection. Additional information related to risk factor counseling would also be helpful.
Plasmodium and Babesia spp.
Plasmodium spp.
Vythilingam I, Lim YA, Venugopalan B, Ngui R, Leong CS, Wong ML, Khaw L, Goh X, Yap N, Sulaiman WY, Jeffery J, Zawiah AG, Nor Aszlina I, Sharma RS, Yee Ling L, Mahmud R. 2014. Plasmodium knowlesi malaria an emerging public health problem in Hulu Selangor, Selangor, Malaysia (2009-2013): epidemiologic and entomologic analysis. Parasit Vectors 7:436. PMID 25223878
A review of 100 Plasmodium knowlesi/Plasmodium malariae cases over a 5-year period in the Hulu Selangor district found these cases to be predominantly seen among young adults (ages 20 to 39 years; 67 cases; 67%); no deaths were reported. In most cases, the individuals worked in agriculture and forestry (51; 51%). Information based on the examination of 535 mosquitoes indicated Anopheles maculatus was the predominant species (49.5%), followed by Anopheles letifer (13.1%) and Anopheles introlatus (11.6%). Only A. introlatus was positive for Plasmodium oocysts. The authors demonstrated that P. knowlesi hotspot areas overlapped with areas where the infected A. introlatus was found, thus strongly supporting the hypothesis that A. introlatus is the vector for P. knowlesiTherefore, A. introlatus has been implicated as a vector of P. knowlesi in Hulu Selangor. These cases of P. knowlesi continue to increase.