Clinical Microbiology Reviews, October 2011

Clinical Microbiology Reviews A Publication of the American Society for Microbiology

VOLUME 24

●

OCTOBER 2011

●

NUMBER 4

CONTENTS/SUMMARIES

Sporothrix schenckii and Sporotrichosis. Mo ˆnica Bastos de Lima Barros, Rodrigo de Almeida Paes, and Armando Oliveira Schubach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

633–654

Summary: Sporotrichosis, which is caused by the dimorphic fungus Sporothrix schenckii, is currently distributed throughout the world, especially in tropical and subtropical zones. Infection generally occurs by traumatic inoculation of soil, plants, and organic matter contaminated with the fungus. Certain leisure and occupational activities, such as floriculture, agriculture, mining, and wood exploitation, are traditionally associated with the mycosis. Zoonotic transmission has been described in isolated cases or in small outbreaks. Since the end of the 1990s there has been an epidemic of sporotrichosis associated with transmission by cats in Rio de Janeiro, Brazil. More than 2,000 human cases and 3,000 animal cases have been reported. In humans, the lesions are usually restricted to the skin, subcutaneous cellular tissue, and adjacent lymphatic vessels. In cats, the disease can evolve with severe clinical manifestations and frequent systemic involvement. The gold standard for sporotrichosis diagnosis is culture. However, serological, histopathological, and molecular approaches have been recently adopted as auxiliary tools for the diagnosis of this mycotic infection. The first-choice treatment for both humans and cats is itraconazole.

Trypanosoma cruzi and Chagas’ Disease in the United States. Caryn Bern, Sonia Kjos, Michael J. Yabsley, and Susan P. Montgomery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

655–681

Summary: Chagas’ disease is caused by the protozoan parasite Trypanosoma cruzi and causes potentially life-threatening disease of the heart and gastrointestinal tract. The southern half of the United States contains enzootic cycles of T. cruzi, involving 11 recognized triatomine vector species. The greatest vector diversity and density occur in the western United States, where woodrats are the most common reservoir; other rodents, raccoons, skunks, and coyotes are also infected with T. cruzi. In the eastern United States, the prevalence of T. cruzi is highest in raccoons, opossums, armadillos, and skunks. A total of 7 autochthonous vector-borne human infections have been reported in Texas, California, Tennessee, and Louisiana; many others are thought to go unrecognized. Nevertheless, most T. cruzi-infected individuals in the United States Continued on following page

Continued from preceding page

are immigrants from areas of endemicity in Latin America. Seven transfusion-associated and 6 organ donor-derived T. cruzi infections have been documented in the United States and Canada. As improved control of vector- and blood-borne T. cruzi transmission decreases the burden in countries where the disease is historically endemic and imported Chagas’ disease is increasingly recognized outside Latin America, the United States can play an important role in addressing the altered epidemiology of Chagas’ disease in the 21st century.

Current Knowledge of Trichosporon spp. and Trichosporonosis. Arnaldo L. Colombo, Ana Carolina B. Padovan, and Guilherme M. Chaves . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

682–700

Summary: Trichosporon spp. are basidiomycetous yeast-like fungi found widely in nature. Clinical isolates are generally related to superficial infections. However, this fungus has been recognized as an opportunistic agent of invasive infections, mostly in cancer patients and those exposed to invasive medical procedures. It is possible that the ability of Trichosporon strains to form biofilms on implanted devices, the presence of glucuronoxylomannan in their cell walls, and the ability to produce proteases and lipases are all factors likely related to the virulence of this genus and therefore may account for the progress of invasive trichosporonosis. Disseminated trichosporonosis has been increasingly reported worldwide and represents a challenge for both diagnosis and species identification. Phenotypic identification methods are useful for Trichosporon sp. screening, but only molecular methods, such as IGS region sequencing, allow the complete identification of Trichosporon isolates at the species level. Methods for the diagnosis of invasive trichosporonosis include PCR-based methods, Luminex xMAP technology, and, more recently, proteomics. Treating patients with trichosporonosis remains a challenge because of limited data on the in vitro and in vivo activities of antifungal drugs against clinically relevant species of the genus. Despite the mentioned limitations, the use of antifungal regimens containing triazoles appears to be the best therapeutic approach.

Clinical Manifestations, Diagnosis, and Treatment of Mycobacterium haemophilum Infections. Jerome A. Lindeboom, Lesla E. S. Bruijnesteijn van Coppenraet, Dick van Soolingen, Jan M. Prins, and Eduard J. Kuijper . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

701–717

Summary: Mycobacterium haemophilum is a slowly growing acid-fast bacillus (AFB) belonging to the group of nontuberculous mycobacteria (NTM) frequently found in environmental habitats, which can colonize and occasionally infect humans and animals. Several findings suggest that water reservoirs are a likely source of M. haemophilum infections. M. haemophilum causes mainly ulcerating skin infections and arthritis in persons who are severely immunocompromised. Disseminated and pulmonary infections occasionally occur. The second at-risk group is otherwise healthy children, who typically develop cervical and perihilar lymphadenitis. A full diagnostic regimen for the optimal detection of M. haemophilum includes acid-fast staining, culturing at two temperatures with iron-supplemented media, and molecular detection. The most preferable molecular assay is a real-time PCR targeting an M. haemophilum-specific internal transcribed spacer (ITS), but another approach is the application of a generic PCR for a mycobacterium-specific fragment with subsequent sequencing to identify M. haemophilum. No standard treatment guidelines are available, but published literature agrees that immunocompromised patients should be treated with multiple antibiotics, tailored to the disease presentation and underlying degree of immune suppression. The outcome of M. haemophilum cervicofacial lymphadenitis in immunocompetent patients favors surgical intervention rather than antibiotic treatment.

Food Animals and Antimicrobials: Impacts on Human Health. Bonnie M. Marshall and Stuart B. Levy. . . . . . . . . . . . . . . . . . . . . . . . . .

718–733

Summary: Antimicrobials are valuable therapeutics whose efficacy is seriously compromised by the emergence and spread of antimicrobial resistance. The provision of antibiotics to food animals encompasses a wide variety of nontherapeutic purposes that include growth promotion. The concern over resistance emergence and spread to people by nontherapeutic use of antimiContinued on following page

Continued from preceding page

crobials has led to conflicted practices and opinions. Considerable evidence supported the removal of nontherapeutic antimicrobials (NTAs) in Europe, based on the “precautionary principle.” Still, concrete scientific evidence of the favorable versus unfavorable consequences of NTAs is not clear to all stakeholders. Substantial data show elevated antibiotic resistance in bacteria associated with animals fed NTAs and their food products. This resistance spreads to other animals and humans—directly by contact and indirectly via the food chain, water, air, and manured and sludge-fertilized soils. Modern genetic techniques are making advances in deciphering the ecological impact of NTAs, but modeling efforts are thwarted by deficits in key knowledge of microbial and antibiotic loads at each stage of the transmission chain. Still, the substantial and expanding volume of evidence reporting animal-to-human spread of resistant bacteria, including that arising from use of NTAs, supports eliminating NTA use in order to reduce the growing environmental load of resistance genes.

Human Metapneumovirus: Lessons Learned over the First Decade. Verena Schildgen, Bernadette van den Hoogen, Ron Fouchier, Ralph A. Tripp, Rene Alvarez, Catherine Manoha, John Williams, and Oliver Schildgen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

734–754

Summary: It has been 10 years since human metapneumovirus (HMPV) was identified as a causative agent of respiratory illness in humans. Since then, numerous studies have contributed to a substantial body of knowledge on many aspects of HMPV. This review summarizes our current knowledge on HMPV, HMPV disease pathogenesis, and disease intervention strategies and identifies a number of areas with key questions to be addressed in the future.

Serratia Infections: from Military Experiments to Current Practice. Steven D. Mahlen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

755–791

Summary: Serratia species, in particular Serratia marcescens, are significant human pathogens. S. marcescens has a long and interesting taxonomic, medical experimentation, military experimentation, and human clinical infection history. The organisms in this genus, particularly S. marcescens, were long thought to be nonpathogenic. Because S. marcescens was thought to be a nonpathogen and is usually red pigmented, the U.S. military conducted experiments that attempted to ascertain the spread of this organism released over large areas. In the process, members of both the public and the military were exposed to S. marcescens, and this was uncovered by the press in the 1970s, leading to U.S. congressional hearings. S. marcescens was found to be a certain human pathogen by the mid-1960s. S. marcescens and S. liquefaciens have been isolated as causative agents of numerous outbreaks and opportunistic infections, and the association of these organisms with point sources such as medical devices and various solutions given to hospitalized patients is striking. Serratia species appear to be common environmental organisms, and this helps to explain the large number of nosocomial infections due to these bacteria. Since many nosocomial infections are caused by multiply antibioticresistant strains of S. marcescens, this increases the danger to hospitalized patients, and hospital personnel should be vigilant in preventing nosocomial outbreaks due to this organism. S. marcescens, and probably other species in the genus, carries several antibiotic resistance determinants and is also capable of acquiring resistance genes. S. marcescens and S. liquefaciens are usually identified well in the clinical laboratory, but the other species are rare enough that laboratory technologists may not recognize them. 16S rRNA gene sequencing may enable better identification of some of the less common Serratia species.

Immunodiagnosis of Tuberculosis: a Dynamic View of Biomarker Discovery. Shajo Kunnath-Velayudhan and Maria Laura Gennaro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

792–805

Summary: Infection with Mycobacterium tuberculosis causes a variety of clinical conditions ranging from life-long asymptomatic infection to overt disease with increasingly severe tissue damage and a heavy bacillary burden. Immune biomarkers should follow the evolution of infection and disease because the host immune response is at the core of protection against disease and tissue damage in M. tuberculosis infection. Moreover, levels of immune markers are Continued on following page

Continued from preceding page

often affected by the antigen load. We review how the clinical spectrum of M. tuberculosis infection correlates with the evolution of granulomatous lesions and how granuloma structural changes are reflected in the peripheral circulation. We also discuss how antigen-specific, peripheral immune responses change during infection and how these changes are associated with the physiology of the tubercle bacillus. We propose that a dynamic approach to immune biomarker research should overcome the challenges of identifying those asymptomatic and symptomatic stages of infection that require antituberculosis treatment. Implementation of such a view requires longitudinal studies and a systems immunology approach leading to multianalyte assays.

CLINICAL MICROBIOLOGY REVIEWS, Oct. 2011, p. 633–654 0893-8512/11/$12.00 doi:10.1128/CMR.00007-11 Copyright © 2011, American Society for Microbiology. All Rights Reserved.

Vol. 24, No. 4

Sporothrix schenckii and Sporotrichosis Mo ˆnica Bastos de Lima Barros1* Rodrigo de Almeida Paes,2 and Armando Oliveira Schubach2 National School of Public Health1 and Evandro Chagas Clinical Research Institute,2 Fiocruz, Rio de Janeiro, Brazil INTRODUCTION .......................................................................................................................................................633 HISTORY.....................................................................................................................................................................634 TAXONOMIC STUDIES ...........................................................................................................................................634 SPOROTHRIX CELL BIOLOGY...............................................................................................................................635 Morphology ..............................................................................................................................................................635 Cell Wall ..................................................................................................................................................................636 S. SCHENCKII GENETIC MATERIAL ...................................................................................................................636 S. SCHENCKII PHYSIOLOGY .................................................................................................................................636 S. SCHENCKII ECOLOGY........................................................................................................................................637 PUTATIVE S. SCHENCKII VIRULENCE FACTORS ...........................................................................................637 Thermotolerance .....................................................................................................................................................638 Melanin ....................................................................................................................................................................638 Adhesion...................................................................................................................................................................638 Ergosterol Peroxide ................................................................................................................................................639 Proteins Related to Virulence ...............................................................................................................................639 IMMUNE RESPONSE IN SPOROTRICHOSIS ...................................................................................................639 Innate Response ......................................................................................................................................................639 Cellular Response ...................................................................................................................................................639 Humoral Response..................................................................................................................................................640 SPOROTRICHOSIS DIAGNOSIS ...........................................................................................................................640 Direct Examination.................................................................................................................................................640 Histopathological Examination.............................................................................................................................641 Culture......................................................................................................................................................................641 Molecular Detection ...............................................................................................................................................641 Sporotrichin Skin Test ...........................................................................................................................................642 Antibody Detection..................................................................................................................................................642 EPIDEMIOLOGY .......................................................................................................................................................643 Geographic Distribution ........................................................................................................................................643 Distribution by Age and Gender...........................................................................................................................643 Transmission and Sources of Infection ...............................................................................................................643 Zoonotic Transmission...........................................................................................................................................644 PATHOGENESIS AND CLINICAL FORMS .........................................................................................................645 Human Disease .......................................................................................................................................................645 Cutaneous forms .................................................................................................................................................645 Mucosal form.......................................................................................................................................................646 Extracutaneous form ..........................................................................................................................................646 Associated signs and symptoms........................................................................................................................646 Differential diagnosis..........................................................................................................................................646 Feline Disease..........................................................................................................................................................646 TREATMENT..............................................................................................................................................................647 Antifungal Susceptibility Tests .............................................................................................................................647 PREVENTION.............................................................................................................................................................649 ACKNOWLEDGMENTS ...........................................................................................................................................649 REFERENCES ............................................................................................................................................................649 matter contaminated with the fungus. Certain leisure and occupational activities, such as floriculture, agriculture, mining, and wood exploitation, are traditionally associated with the mycosis. Zoonotic transmission has been described in isolated cases or in small outbreaks. At present, veterinarians, technicians, caretakers, and owners of cats with sporotrichosis are regarded as a new risk category for the acquisition of the disease. The lesions are usually restricted to the skin, subcutaneous cellular tissue, and adjacent lymphatic vessels. Eventually, this fungus can disseminate to other organs, and alternatively, on rare occasions, inhalation of conidia may lead to a

INTRODUCTION Sporotrichosis, caused by the dimorphic fungus Sporothrix schenckii, is currently distributed throughout the world, especially in tropical and subtropical zones. Infection generally occurs by traumatic inoculation of soil, plants, and organic

* Corresponding author. Mailing address: Fundac¸˜ao Oswaldo Cruz/ Fiocruz, Av. Brasil, 4365, Manguinhos, Rio de Janeiro CEP 22045-900, Brazil. Phone: 55 21 2598 2518. Fax: 55 21 3415 2086. E-mail: [email protected]. 633

634

BARROS ET AL.

CLIN. MICROBIOL. REV.

systemic disease. Several factors, such as inoculum load, immune status of the host, virulence of the inoculated strain, and depth of traumatic inoculation, influence the different clinical forms of sporotrichosis. The gold standard for sporotrichosis detection is culture; however, serological, histopathological, and molecular approaches have been recently adopted as auxiliary tools for the diagnosis of this mycotic infection. HISTORY Sporothrix schenckii was isolated for the first time in 1896 by Benjamin Schenck, a medical student at the Johns Hopkins Hospital in Baltimore, MD, from a 36-year-old male patient presenting lesions on the right hand and arm. This isolate, from the patient abscess, was then studied by the mycologist Erwin Smith, who concluded that the fungus belonged to the genus Sporotrichum (217). Previously, Linck in 1809 and Lutz in 1889 referred to some possible sporotrichosis cases, but the isolation of the fungus by these authors for case definitions was not possible (126). The second undeniable sporotrichosis case was described in 1900 by Hektoen and Perkins, also in the United States (Chicago, IL). This was the case of a boy who suffered an injury with a hammer hitting his finger, with the lesion presenting spontaneous regression. These investigators gave the sporotrichosis agent its current denomination, Sporothrix schenckii (95). Later, this fungus was erroneously included in the genus Sporotrichum, which comprises basidiomycetous fungi which are neither dimorphic nor pathogenic for humans or other animals (216). This erroneous nomenclature remained until 1962, when Carmichael recognized differences in the conidiations of members of the genus Sporotrichum and isolates from sporotrichosis cases (37). In 1903, Sabouraud suggested to Beurmann and Gougerot the use of potassium iodine for the treatment of sporotrichosis, which was a common disease in France during the beginning of the 20th century (126). This has hitherto been a satisfactory therapy for sporotrichosis, although no randomized, doubleblind, placebo-controlled trials have ever been conducted (267). The first reported case of natural animal infection was described in 1907 by Lutz and Splendore in rats from Brazil (141). The possibility of human infection by bites from these rats was considered (186). Also in Brazil, in 1908, Splendore reported the detection of asteroid bodies around Sporothrix yeast cells, which offer a very useful tool for sporotrichosis diagnosis in histological examinations (126, 196). TAXONOMIC STUDIES Sporothrix schenckii belongs to the kingdom Fungi and is a eukaryotic organism that is without mobility and heterotrophic and presents chitin on its cell wall. For several years, this fungus was included in division Eumycota, subdivision Deuteromycotina, class Hyphomycetes, order Moniliales, and family Moniliaceae (128). After a substantial fungal taxonomy revision by Guarro and coworkers, this fungus was characterized in division Ascomycota, class Pyrenomycetes, order Ophiostomatales, and family Ophiostomataceae (84).

The sexual form of S. schenckii is as yet unknown. However, there is substantial molecular evidence that this fungus undergoes recombination in nature (163). Nevertheless, some studies imply that S. schenckii in an ascomycete, since it presents a simple septum, with Woronin bodies (237) and three chitin synthase genes (44). Molecular analyses of the 18S region of the ribosomal DNA indicate that the sexual form of S. schenckii could be Ophiostoma stenoceras (22). On the other hand, morphological and physiological studies exhibit consistent differences between these two species. O. stenoceras is unable to produce dematiaceous conidia, as does S. schenckii. Also, S. schenckii does not produce perithecium on malt, rice, or potato media, as is observed for isolates of O. stenoceras (60, 181). Differences are also apparent when these species are inoculated in mice. S. schenckii can be found in several tissues from all infected mice after intravenous inoculation, and O. stenoceras is detected in certain organs from some infected animals (59). These observations lead to the conclusion that the O. stenoceras anamorph and S. schenckii are different species. Meanwhile, other molecular studies (56, 97), together with work by Berbee and Taylor (22), reinforce that the S. schenckii teleomorph is classified in the genus Ophiostoma. Berbee and Taylor highlight that S. schenckii belongs to the pyrenomycete lineage, lacking forcible ascospore discharge (22). Recently, Marimon and coworkers (150), on the basis of phenotypic and genotypic analyses, suggested that S. schenckii should not be considered the only species that causes sporotrichosis, and based on macroscopic characteristics, sucrose and raffinose assimilation, ability to grow at 37°C, and the nuclear calmodulin gene sequence, they described four new species in the Sporothrix complex: (i) S. globosa, a fungus distributed worldwide (145, 180); (ii) S. brasiliensis, the species related to the zoonotic epidemic of sporotrichosis in Rio de Janeiro, Brazil (150, 179); (iii) S. mexicana, limited to Mexico (150); and (iv) S. luriei, formerly S. schenckii var. luriei (151), differing from S. schenckii mainly in the tissue form by the production of large, often septate budding cells unable to assimilate creatinine or creatine (53). On the other hand, other authors support its separation by rRNA internal transcribed spacer (ITS) sequence data (54). Another species, S. cyanescens has been isolated from blood and skin samples from human patients, but pathogenicity studies conclude that, although this fungus can grow at 37°C, it is avirulent (233). Figure 1 presents a key to differentiate species within the S. schenckii complex (151). Recently, de Meyer and collaborators (56) described three other environmental Sporothrix species, S. stylites, S. humicola, and S. lignivora. The first two species differ from S. schenckii by the inability to produce melanized conidia and the consequent nondarkening of colonies with age. S. lignivora has distinctive conidia that do not match in size and shape those of other Sporothrix or Ophiostoma species. It is interesting to note that isolates classified as S. humicola were previously referred to as environmental isolates of S. schenckii. In their study, the authors concluded that ␤-tubulin sequence analysis is strongly recommended for taxonomic studies of Sporothrix species isolated from the environment.

VOL. 24, 2011

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

635

FIG. 1. Identification key for Sporothrix species of clinical interest, based on morphological and phenotypic tests described by Marimon and collaborators (152). PDA, potato dextrose agar; CMA, corn meal agar.

SPOROTHRIX CELL BIOLOGY Morphology Sporothrix schenckii is a dimorphic fungus. In its saprophytic stage or when cultured at 25°C, it assumes a filamentous form, composed of hyaline, septate hyphae 1 to 2 ␮m wide, with conidiogenous cells arising from undifferentiated hyphae forming conidia in groups on small, clustered denticles. These onecelled conidia are tear shaped to clavate (53) and do not yield chains (233). Often, hyaline or brown thick-walled conidia arise beside the hyphae. The dark cell walls of the conidia distinguish S. schenckii from other, nonpathogenic Sporothrix species (56, 242). Macroscopically, filamentous colonies in media such as malt extract agar or potato dextrose agar (Fig. 2) are often smooth and wrinkled, white to creamy at first and then turning brown to black after a few days (126, 170, 196). Some strains, however, have the ability to form dark colonies from the beginning of growth (5). The S. schenckii colonies never become cottony or floccose (126). This fungus is evident in both human and animal tissues as budding yeasts. Yeastlike cells can be observed in various sizes and shapes. They may be round to oval, with 2- to 6-␮m diameters, and usually have elongated, cigar-shaped buds on a narrow base. Macroscopically, yeast colonies (Fig. 2) are smooth, tan, or cream colored (130). Some molecular aspects implicated in proliferation and maintenance of this morphological form of S. schenckii involve calcium/calmodulin-dependent protein kinases (260) and a signaling pathway involving the interaction between a cytosolic phospholipase and protein G. Studies have proven that this pathway is necessary for the reentry of S. schenckii yeast cells into the budding cycle, suggesting its function in the control of dimorphism in this fungus and for the maintenance of the yeast form (259). The transition from mold to yeast form in S. schenckii can be attained by culturing mycelia or conidia on rich culture media such as brain heart infusion agar at 35 to 37°C (170). Some strains, especially those related to the S. globosa species, may

require lower conversion temperatures, since they do not grow well at 37°C (150). Although rich media are required for the mycelium-to-yeast transition, S. schenckii yeast cells can be maintained at 37°C in other media, such as Sabouraud dextrose agar. This transition process also occurs after patients are infected with filamentous S. schenckii. Morphological transformation at the ultrastructural level occurs by direct formation of budlike structures at the tips and along the hyphae together with oidial cell formation after septation of the hyphae, without conspicuous alterations of the cytoplasmic content of parent mycelial cell. There is no direct budding of yeast from conidiospores (77).

FIG. 2. Cultures of pus from lesions of S. schenckii-infected patients. Most strains become visible after 4 days of growth on Sabouraud dextrose agar, presenting no visible dark pigment at this stage (tube at left), whereas others are melanized since the beginning of growth (tube at center). When transferred to brain heart infusion agar and cultured at 37°C, strains undergo dimorphism, presenting creamy white to tan yeast colonies after 7 days of growth (tube at right).

636

BARROS ET AL.

CLIN. MICROBIOL. REV.

The beginning of the yeast-to-mycelium transition in S. schenckii is a process regulated by calcium, which induces both RNA and protein synthesis on the yeast cell (199). A prerequisite for this transition process is a nuclear division; afterwards, a germ tube is originated from the parental yeast cell and a septum is formed at mother cell-germ tube formation (25). It is interesting to note that yeast cells can be maintained at 25°C if cultured in liquid media with glucose and with the pH around 7.2 (195, 260). Cell Wall Like other fungi, S. schenckii has a cell wall surrounding the plasma membrane in both the mycelial and yeast forms. There are characteristic differences in cell wall thickness between conidia, yeast forms, and filaments, as well variations in plasma membrane invaginations among these three morphological forms of the fungus. It has been shown by freeze fracturing studies that in conidia invaginations are short and abundant and in yeast forms they are scarce and longer, while the plasma membrane of the S. schenckii hyphae is smooth, without invaginations (244). The fungal cell wall is rigid as well as complex, and recently it has been shown that S. schenckii produces vesicles that are probably related to the transfer of periplasmic molecules and pigment-like structures from the plasma membrane to the extracellular space, since in contrast to the case for prokaryotic organisms, in eukaryotic cells there is vesicular traffic of molecules to the plasma membrane (3, 197). The chemical structures of fungal cell wall polysaccharides and glycoproteins have been studied basically because of the knowledge of the antigenic structures of human pathogens (193). A peptide-rhamnomannan was isolated from the yeast S. schenckii cell wall, where there were the polysaccharides D-mannose (50%) and L-rhamnose (33%), small amounts of galactose (1%), and about 16% peptides (138). Comparative studies of mycelial and yeast S. schenckii cell walls showed little difference in the glycosidic components. The cell wall composition of the mycelial phase included large amounts of lipids and protein and a lower concentration of mannose (193). The cell wall composition in S. schenckii conidial cells can also be affected by the time of culture, with a decrease in the rhamnose molar ratio and an increase in the mannose molar ratio (67). The yeast cell wall of S. schenckii also contain granules of melanin (250) and proteins involved in adherence (135, 206), which contribute to fungal virulence. Of particular interest is a glycoprotein of 70 kDa isolated from the cell wall of the S. schenckii yeast phase. The purified glycopeptide has a pI of 4.1, and about 5.7% of its molecular mass is composed of N-linked glycans, with no evidence for O-linked oligosaccharides in this molecule. This glycoprotein has a uniform distribution on the fungal cell surface and participates in adhesion to the dermal extracellular matrix (206). S. SCHENCKII GENETIC MATERIAL Little is known about the S. schenckii genomic composition because this fungus is not amenable to genetic analysis based on meiotic segregation (256). Studies on the genomic DNA base composition rendered an average guanine and cytosine

content of about 54.7 mol%, with the DNA showing a low degree of hybridization with O. stenoceras DNA, supporting the supposition that this fungus does not represent the sexual state of S. schenckii. However, 75% hybridization was observed with Ophiostoma minus DNA (159). More recently, Tateishi and coworkers (249), karyotyping eight strains isolated from patients in Japan, concluded that S. schenckii possesses six to eight chromosomes of 460 to 6,200 kb, with a total genome size of approximately 28 Mbp. Another study with strains from a different geographical origin predicted a 45-Mbp genome size for S. schenckii (256). Perhaps these differences are related to either the different species recently described (150) or to underestimations in the methods adopted for genome size determination. Also, it has been reported that S. schenckii is a diploid organism, bearing around 50 fg DNA per cell, in both the filamentous and yeast phases. On the other hand, aneuploidy, a state in which most of the chromosomes are disomic, cannot be excluded (256). It is interesting to note that diploidy is essential for thermal dimorphism in Cryptococcus neoformans, and similarities in life cycle between this fungus and other dimorphic fungi, including S. schenckii, may occur (232). Studies on identification, typing, and epidemiology of sporotrichosis are usually based on mitochondrial DNA (mtDNA) analysis of restriction length polymorphisms (RFLP) with the restriction enzyme HaeIII. Initially, 24 mtDNA types were cited (136), and more recently types 25 to 30 (168) and 31 to 32 (103) were introduced. These analyses have been adopted in several studies with S. schenckii strains from different geographical origins and also environmental isolates (13, 103, 104, 243, 266). S. SCHENCKII PHYSIOLOGY Even though sporotrichosis is a disease distributed worldwide, there are only a few studies regarding the physiological characteristics of its agent. In general, the optimal temperature for S. schenckii growth is around 30 to 37°C, with growth of all strains being impeded at 40°C (79, 150). Although S. schenckii is able to grow at 35 to 37°C, some growth inhibition is observed compared to that at 28°C. Moreover, this inhibition appears to be geographically related (163). Several carbohydrates can be assimilated by S. schenckii, such as glucose, fructose, mannose, and cellobiose (79, 200). However, there is some variability in assimilation of sucrose, arabinose, starch, raffinose, and ribitol (79, 150). Starch assimilation is also affected by fungal preservation under some storage methodologies, such as the Castellani method (160). The carbohydrate concentration available during S. schenckii growth modulates melanin synthesis by this fungus (Fig. 3), enhancing pigment formation in a glucose concentration-dependent manner (5). This fungus is not able to ferment any carbohydrate (53). Some physiological differences between the two different S. schenckii morphologies may be observed. Mycelial-phase S. schenckii can grow well at pHs of around 3.0 to 11.5, but yeast cells can grow only within the pH range of 3.0 to 8.5. The yeast phase is also more osmotolerant (30%) than the mycelial phase (20%), as is true for halophila. The mycelial phase withstands growth in 7% NaCl, but the yeast-phase S. schenckii can grow well in 11% NaCl (79).

VOL. 24, 2011

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

FIG. 3. Glucose concentration-dependent increase of melanin synthesis. The concentration of glucose (%, wt/vol) in each culture is indicated by the numbers on the agar plates. S. brasiliensis strain 17307, grown at 22°C (A), and S. schenckii strain 23250, grown at 37°C (B), are representative isolates showing enhancement of melanization with increasing glucose amounts.

S. schenckii is able to split urea (79, 150, 233), perform reductive iron acquisition with secreted extracellular enzymes (270), and tolerate cycloheximide at 0.25% (150). Thiamine is required for fungal growth (60, 101). S. SCHENCKII ECOLOGY S. schenckii often enters the host through traumatic implantation (126, 129, 170, 196). In nature, the fungus has been found to live as a saprophyte on living and decaying vegetation, animal excreta, and soil (118, 126, 156, 162). Organic material in soil is fundamental for mycelium development. The fungus thrives in soil plentiful in cellulose, with a pH range from 3.5 to 9.4 and a temperature of 31°C (177). The relative humidity cannot be below 92% (126). It has been proposed that the armadillo Dasypus septemcinctus may be a reservoir of S. schenckii, since armadillo hunting was reported by several patients with sporotrichosis in Uruguay (143). In fact, this armadillo harbors S. schenckii in neither its intestine nor its epidermis, but the fungus can be found on the dry grass used by these animals for nesting (126). Here we find an apparent incongruity between the organisms needing high humidity for growth (126) yet being found on dry grass of armadillo nests. Probably these differences are geographically related, like growth inhibition at high temperatures (163). Moreover, we cannot discard the hypothesis that different species within the S. schenckii complex (150, 151) have different humidity requirements for growth. There is another armadillo species, Dasypus novemcinctus, which is susceptible to systemic, fatal sporotrichosis (111, 264). Other animals related to S. schenckii transmission are parrots, rodents, cats, dogs, squirrels, horses, and birds (126, 212). Moreover, S. schenckii was also isolated from aquatic animals, primarily fish and dolphins (91, 164), as well as from insects that had been in direct contact with the fungus (126). Some authors have reported sporotrichosis cases due to mosquito bites (129).

637

It has been confirmed that S. schenckii is associated with plants. Sphagnum moss, rose thorns, and hay are especially recognized to harbor this pathogenic fungus (64, 71, 156). However, S. schenckii does not have the potential to be a plant pathogen, probably because extracts from several plants have antifungal activity against S. schenckii (81, 153, 154, 213). In fact, it has been described that when S. schenckii is inoculated in living or dead sphagnum moss, the fungal cell population proliferates in the moist dead plants but not in live moss (271), suggesting that plants have some mechanism to control S. schenckii overgrowth. There has been some reports about S. schenckii isolation from food (1, 117). Nevertheless, this fungus does not appear to have the potential to cause food-borne infection (116). There are some reports on the isolation of S. schenckii from environmental sources. Methods include direct isolation of the fungus by plating the supernatants of samples suspended in water or physiological saline solution with antibiotics in Mycosel agar medium or by inoculation of this suspension in susceptible mice, such as BALB/c, with further culture of spleens, livers, and lungs of the infected animals (60, 162). Direct isolation appears to be more effective to obtain S. schenckii from environmental samples. However, colonies obtained after mouse inoculation have been free of microbial contamination (162). PUTATIVE S. SCHENCKII VIRULENCE FACTORS We can define a virulence factor as a feature of a microorganism that allows or enhances microbial growth in the host. To study and characterize these factors, is necessary compare the microbe-host interactions of an isolate that expresses the suspected factor and a mutant isolate that has lost the ability to express it, which can be attained by induced mutagenesis through molecular strategies. If differences in the infections caused by these different isolates are noted, it is imperative to make the mutant isolate recover the ability to express the studied factor and check whether or not the organism then regains the capacity to cause infection similar to that of the parental wild-type strain (100). Discovery of the origin of microbial virulence has been the main goal of several studies. In general, the most accepted theory is that with microbial interactions with other organisms present in the natural habitat of the pathogen, the microorganisms acquire survival strategies tending to a higher virulence when they accidentally find an animal host. For instance, these microorganisms, in the mammalian host, usually have the ability to form biofilms and mechanisms to acquire iron and produce proteolytic enzymes that will lead to higher virulence (40). Regarding this theory, Steenbergen and coworkers (240) suggested that the origin of virulence in S. schenckii should be related to the intermicrobial interactions in its environment. The authors demonstrated that when ingested by Acanthamoeba castellanii, a soil amoeba, S. schenckii yeast cells are able to survive within the protozoan, kill it, and use it as nutrient. This behavior is not shared by pathogenic fungi that do not have the soil as habitat, such as Candida albicans, or by fungi that are not primarily pathogenic, such as Saccharomyces cerevisiae. On the other hand, other dimorphic fungal patho-

638

BARROS ET AL.

CLIN. MICROBIOL. REV.

FIG. 4. Melanin ghosts of S. schenckii 18782 strain under several culture conditions. (A) Cultures on minimal medium at 25°C yield melanin ghosts only from dematiaceous conidia. (B) When L-DOPA is added to minimal medium, both hyphae and conidia are melanized. (C) Yeast S. schenckii cells can also produce melanin in culture medium free of phenolic compounds or when L-DOPA is added. Bars, 10 ␮m.

gens, such as Histoplasma capsulatum and Blastomyces dermatitidis (240) as well as Cryptococcus gattii, a highly virulent yeast pathogen (148), have the same behavior as S. schenckii when in contact with A. castellanii. Little is known about S. schenckii virulence factors due to the lack of studies in this field, in part because S. schenckii is not responsive to genetic analysis. However, some putative virulence factors have appeared from some investigations. Thermotolerance One of the putative S. schenckii virulence factors, which is also a virulence factor of other pathogenic fungi, is thermotolerance (40). In fact, isolates able to grow at 35°C but not at 37°C are incapable of causing lymphatic sporotrichosis and produce fixed cutaneous lesions instead. The fungi isolated from lymphatic, disseminated and extracutaneous lesions show tolerance and growth at 37°C (126). Results from a more recent study demonstrate that S. schenckii isolates from Colombia, where most patients are affected with fixed cutaneous sporotrichosis, exhibit high growth inhibition at 35 and 37°C, in contrast to isolates from Mexico and Guatemala, where lymphatic sporotrichosis prevails (163). In an in vivo mouse assay, it has been shown that if the mice have their feet warmed in cages with heat on the floor, progression of sporotrichosis is reduced compared to that in control infected mice maintained in regular cages. Isolates from pulmonary lesions, however, replicate regularly in both groups of mice (246). Melanin Both morphological stages of S. schenckii have the ability to synthesize melanin. This is an insoluble compound highly related to virulence in several fungi (106). Melanin production in S. schenckii dematiaceous conidia occurs through the 1,8-dihydroxynaphthalene (DHN) pentaketide pathway (202). Macroscopically, only the mycelial phase of the fungus is melanized. However, melanin production in yeast cells was demonstrated in vitro during infection (171). Recently, it has been demonstrated that S. schenckii can also produce melanin using phenolic compounds such as 3,4-dihydroxy-L-phenylalanine (L-DOPA) as a substrate both in filamentous and yeast forms (5). It is interesting to note that only conidia can be melanized by the DHN pathway, but if L-DOPA is present, hyphae can be melanized as well (Fig. 4). Since S. schenckii is a soil-accommodated fungus that does

not require host parasitism to complete its life cycle, fungal melanization must be also important against unfavorable environmental conditions, since mycelium is the fungal form encountered in nature (171). In vitro studies indicate that melanization in S. schenckii is controlled by several factors, such as temperature, pH, and nutrient conditions (5). Moreover, similar culture media from different suppliers can yield differences in melanization within a single S. schenckii strain (250). It has been shown that conidial melanization enhances S. schenckii resistance to macrophage phagocytosis, allowing the first steps of infection, since these structures usually are the fungal infective particles (202). Corroborating this hypothesis, it has been demonstrated by molecular typing of an S. schenckii strain isolated from a laboratory worker who had handled a pigmented strain and an albino strain of S. schenckii that the isolate from the patient had the same genotypic profile as the dematiaceous strain (48). Melanization also has a role in the pathogenesis of cutaneous sporotrichosis, since pigmented isolates had a greater invasive ability than the albino mutant strain in an experimental rat model of sporotrichosis. The albino strain also was restricted to the core of the granuloma. In addition, the melanized strain promoted the formation of multifocal granulomas (145). Some S. schenckii melanization in vivo has been previously described, such as a weak brown halo on the yeast S. schenckii cell wall when infected sporotrichosis tissues are stained with Fontana-Masson stain, a technique initially developed to demonstrate melanin in C. neoformans (127). This hypothesis has now been confirmed by detection of Sporothrix melanin ghosts in tissues from infected animals as well as by detection of antimelanin antibody in sera from patients with sporotrichosis (5, 171). Since melanization decreases the susceptibilities of H. capsulatum and C. neoformans to amphotericin B and caspofungin (261), melanin pigment in S. schenckii may hamper treatment in some sporotrichosis cases, especially in cases of extracutaneous disease or in patients infected with human immunodeficiency virus (HIV) (171). However, there have been no studies confirming this hypothesis. Adhesion Primary adhesion to endothelial and epithelial cells as well as on extracellular matrix components is essential to an effective invasion of host tissues by pathogens. Both conidia and yeast cells from S. schenckii are able to recognize three impor-

VOL. 24, 2011

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

tant glycoproteins from the extracellular matrix: fibronectin, laminin, and type II collagen (134, 135). Some studies have demonstrated that the fungus has integrins or adhesin lectinlike molecules that recognize human fibronectin at several points on the molecule (135). The fibronectin adhesins are located on the surface of yeast cells, and their expression is related to fungal virulence (251). It is also known that these fibronectin receptors are different from laminin receptors (133). These receptors are present on both hyphae and fungal yeasts, although yeasts have a greater ability to bind to the extracellular matrix. The existence of these adhesins would favor adherence to host tissues and fungal dissemination throughout the body (133). Expression of these molecules in S. schenckii is probably related to virulence, since their preferential expression is in the parasitic rather than the saprophytic form of the fungus. Recently a 70-kDa glycoprotein from an S. schenckii isolate was described, and its participation in adhesion to the dermal extracellular matrix was demonstrated (206). This fungal pathogen is also able to interact in vitro with human endothelial cells, which can internalize fungal yeast cells without injury or decreased viability. Moreover, the fungus can also cross the intercellular space. Both processes facilitate fungus bloodstream penetration and consequent hematogenous dissemination (69). Transendothelium migration occurs through a paracellular route involving extracellular matrix proteins, in a process mediated by transforming growth factor ␤1 (TGF-␤1) (70). Although the endothelial proteins responsible for this interaction have been characterized, fungal proteins needed for recognition of and adhesion to these cells are unknown, and their part in fungal virulence requires clarification. Ergosterol Peroxide Sgarbi and coworkers, analyzing lipids from S. schenckii through spectroscopic methods, have identified ergosterol peroxide from S. schenckii yeast cells. This compound can be converted to ergosterol when in contact with an enzyme extract from the fungus. The ergosterol peroxide, found in a pathogenic fungus for the first time in S. schenckii, is formed as a protective mechanism to evade reactive oxygen species during phagocytosis and may also represent a virulence factor. Apparently, however, survival of virulent S. schenckii yeast cells after phagocytosis of polymorphonuclear host cells relies on other detoxification strategies besides the one leading to ergosterol peroxide synthesis (32, 231). Proteins Related to Virulence Roles of diverse proteins in the virulence of different fungal pathogens have been described. For instance, the Paracoccidioides brasiliensis immunodominant antigen, a glycoprotein of 43 kDa, is the molecule responsible for laminin and fibronectin recognition and binding, which increase fungal virulence (158, 262). Calcium binding proteins are important in H. capsulatum virulence, enabling acquisition of this ion in environments with calcium limitations (265). A series of virulence-related proteins, such as different adhesins, have been described for Aspergillus fumigatus, including a 30-kDa hemolysin contain-

639

ing several proteases that favor pulmonary colonization and destruction of effective humoral molecules and a 350-kDa catalase needed for phagocytosis survival (131). However, the function of different S. schenckii proteins in virulence is still unclear. It is believed that acid phosphatases act on fungusmacrophage interactions, although no definite evidence to support this theory exists (100). Peptido-rhamnomannans of the fungal cell wall cause depression of immune response until the sixth week of infection and may act as a virulence factor (34). An antigenic preparation from the S. schenckii yeast phase shows proteolytic activity against different human IgG subclasses, suggesting that some secreted proteins may interfere with the immune response of the host (204). Due to the lack of information, characterization of S. schenckii proteins and determination of new virulence factors are imperative for a better understanding of sporotrichosis pathogenesis. IMMUNE RESPONSE IN SPOROTRICHOSIS The virulence of S. schenckii is one of the factors thought to play a role in the development of sporotrichosis (32), but there are discordant results concerning disease evolution in experimental sporotrichosis with S. schenckii clinical isolates from cutaneous and disseminated infection (29, 176), indicating that host immune responses also substantially participate in the progress of sporotrichosis (32). The immunological mechanisms involved in prevention and control of S. schenckii infections are still not very well understood. However, they probably include both humoral and cellular responses (32, 33, 147), which appear to be triggered by distinct antigens. Surface cell antigens, especially some lipids, inhibit the phagocytosis process, while the humoral response is induced by secreted fungal proteins, the exoantigens, which are not involved in the cellular response (35). The innate immune response also plays a role in the pathogenesis of sporotrichosis (32). Innate Response A complement system can be activated by S. schenckii, especially the alternate pathway, although classic complement activation cannot be excluded. Complement activation may support fungal yeast cell phagocytosis by C3b component deposition on the fungal cell wall. The membrane attack complex also contribute to fungal cell lysis (230, 255). Recent studies have emphasized the importance of Toll-like receptor 4 (TLR4) in sporotrichosis. TLR4, also designated CD284, is an important molecule involved in the activation of the innate immune system that, in sporotrichosis, is able to recognize molecules within a lipid extract from the yeast form of the fungus. This interaction leads to the induction of an oxidative burst against the fungus (32). Cellular Response Acquired immunity against the fungus requires the action of activated macrophages. They can be activated during sporotrichosis by CD4 T lymphocytes, which release gamma interferon (IFN-␥), a strong macrophage activator (247), and by other antigen-presenting cells, establishing a link between innate and

640

CLIN. MICROBIOL. REV.

BARROS ET AL.

adaptive immune responses (32). Tumor necrosis factor alpha (TNF-␣), a cytokine that acts on activated macrophages to produce nitric oxide (35), an antioxidant product presenting a high cytotoxic effect against S. schenckii (66), is produced upon incipient and terminal infection, hopefully providing total resolution (147). Although nitric oxide is a fungicidal molecule, this compound may be implicated in immunosuppression in vivo, because high levels of TNF-␣ and NO released after yeast dissemination into the tissues lead to the induction of molecules suppressing T cell responses, such as interleukin10 (IL10), FasL, and CTLA-4. This deleterious effect of NO occurs just upon initial infection, becoming crucial some time after fungal inoculation (68). In fact, TNF-␣ production drastically subsides at 4 to 6 weeks after experimental infection, inducing the fungus to reproduce and infect host tissues. The opposite situation occurs 2 months after infection, when the levels of IL-1 and TNF-␣ increase, favoring fungal elimination (36). After the phagocytosis of S. schenckii conidia and yeast cells, monocytes and macrophages are also strongly induced to produce reactive oxygen species (202). These reactive species, especially superoxide anion and its oxidative reactive metabolites, which are also produced by neutrophils, are involved in fungistatic and fungicidal responses, and their absence is related to a higher lethality in mouse experimental infections (108). Therefore, the Th1 response is of great importance in sporotrichosis pathogenesis, acting as the key factor in controlling fungal infection and with its differential activation leading to varied clinical manifestations of the disease (258). Similar observations on the activation of Th1 cells have led to them being seen as being responsible for different clinical manifestations in other cutaneous infectious diseases, such as leishmaniasis (38).

Humoral Response The humoral immune response is driven by IL-4 produced by Th2 cells. In experimental sporotrichosis, IL-4 release is enhanced at 5 to 6 weeks after infection (147), suggesting the participation of the humoral immune response only in the advanced stages of sporotrichosis (32). Antibodies may have some effect on S. schenckii development, since a monoclonal antibody against a glycolipid antigen is able to hinder S. schenckii growth and differentiation in vitro (254). A monoclonal antibody against the 70-kDa adhesin is also protective in murine model of sporotrichosis (174). Nonetheless, little is known about antibodies elicited during the course of sporotrichosis. It has been described that mice infected with S. schenckii are able to produce specific IgG1 and IgG3 antibodies against a 70-kDa fungal protein during experimental infection, with these antibodies perhaps being related to fungal elimination in these organisms (173). During human sporotrichosis, our group has demonstrated the production of IgG, IgM, and IgA antibodies against mycelial-phase S. schenckii exoantigens. Nevertheless, since patients with different clinical forms of sporotrichosis produce similar amounts of these antibodies, we believe that the humoral immune response against proteins secreted by S. schenckii does not play a role in sporotrichosis pathogenesis (6).

FIG. 5. Direct examination of clinical specimens for diagnosis of sporotrichosis. (A) KOH mount of a tissue fragment from a cat with sporotrichosis, showing cigar-shaped (arrow) and budding (dashed arrow) S. schenckii yeast cells. Note the high fungal burden in the specimen. (B) Direct examination (10% KOH) of the pus from a lesion of a human patient with sporotrichosis, showing nonspecific budding yeast cells. Bars, 10 ␮m. (Courtesy of Rosani Santos Reis.)

SPOROTRICHOSIS DIAGNOSIS Sporotrichosis can be diagnosed through a correlation of clinical, epidemiological, and laboratory data. Laboratory analysis for the determination of sporotrichosis includes direct examination of specimens such as tissue biopsy specimens or pus from lesions. In case of disseminated infections, other specimens, such as sputum, urine, blood, and cerebrospinal and synovial fluids, can be analyzed, depending on the affected organs. Direct Examination Direct examination of specimens is usually conducted with 10% potassium hydroxide in order to observe parasitic budding yeast cells. These yeasts are small (2 to 6 ␮m in diameter) and scarce and consequently are difficult to detect upon direct examination of specimens collected from humans. However, when the same test is performed with samples collected from infected cats, due to the high fungal burden in these animals, yeast cells can be easily found, even at a magnification of ⫻400 (Fig. 5). Fluorescent-antibody staining can help in the observation of yeast forms of S. schenckii; however, this is not a technique that is readily available in most laboratories (130), especially in underdeveloped countries. When the Gram stain is used on the clinical material, yeast cells appear positively stained, sometimes within giant cells or polymorphonuclear lymphocytes (129). For the detection of S. schenckii, some authors recommend Giemsa stain after 10 to 15 dilutions of

VOL. 24, 2011

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

pus in physiological solution (12, 129). These staining procedures also lack sensitivity. Observation of yeast cells through direct examination, however, is not conclusive for sporotrichosis diagnosis. The characteristic “cigar-shaped” buds (2 by 3 to 3 by 10 ␮m) are not always witnessed. Moreover, yeast cells of H. capsulatum and Candida glabrata may be misidentified as S. schenckii (130). Direct examination of pus obtained from lesions of patients with sporotrichosis, without potassium hydroxide, also permits the detection of asteroid bodies. These structures were confirmed in 43.75% of patients, and the sensitivity of the examination can be enhanced (up to 93.75%) if the initial pus is discarded and new samples are collected more deeply. This can help to initiate specific treatment before the results of the culture examinations are available (78).

641

tive of a localized immunological host response to antigens of diverse infectious organisms, including fungi, bacteria, and other parasites. It appears as radiating homogenous, refractile, eosinophilic clublike material surrounding a central eosinophilic focus (130). There are several reports concerning asteroid bodies, the Splendore-Hoeppli reaction in sporotrichosis, in histopathological tissue sections from sporotrichosis patients, ranging in positivity from 20 to 66% (78). Other authors, however, report an absence of this structure in analyzed samples (187). Yeast cells remain viable inside the asteroid bodies, which present IgG and IgM from the host on the spikes of the radiated crowns, suggesting that asteroid bodies are resistance structures which use immune molecules of the host to advantage the yeasts (203). Culture

Histopathological Examination Although S. schenckii may be seen in tissue with the routinely used hematoxylin and eosin (H&E) stain, other special stains such as Gomori methenamine silver (GMS) or periodic acid-Schiff (PAS) stain can be employed to enhance fungal detection (130, 170). Fontana-Masson staining is negative (54). Atypical S. schenckii cells can appear spherical and surrounded by a PAS-positive capsule, resembling Cryptococcus cells (126). Once again, parasitic cells of S. schenckii are difficult to visualize due to the paucity of yeasts in lesions from humans (170) or other animals such as dogs (57). Tissue reaction must be also evaluated in histopathological examinations from patients with sporotrichosis. S. schenckii usually causes a mixed suppurative and granulomatous inflammatory reaction in the dermis and subcutaneous tissue, frequently accompanied by microabscess and fibrosis. Cutaneous infections may also exhibit hyperkeratosis, parakeratosis, and pseudoepitheliomatous hyperplasia (130). Foreign bodies of vegetal origin related to the traumatic inoculation of the agent may also be encountered (182). Besides intact polymorphonuclear cells, granulomas in sporotrichosis usually contain cellular debris, caseous material, giant and epithelioid cell lymphocytes, plasmocytes, and fibroblasts as well as S. schenckii yeast cells within phagocytic cells or in the extracellular medium (129). Miranda and collaborators (167) reported that in dogs with sporotrichosis, lesions present well-formed granulomata, with marked neutrophil infiltration. The peripheral infiltrate often is devoid of lymphocytes and macrophages. Taken together, this information enables the differentiation of sporotrichosis and leishmaniasis in these animals. Some histopathological alterations, such as presence or predominance of epithelioid granulomas, presence of foreign body granulomas, predominance of lymphocytes, presence or predominance of caseous necrosis, and predominance of fibrinoid necrosis and fibrosis, are related to the lack of observation of the fungus in tissue sections from human patients. When the fungus is not in evidence, suppurative granulomas, neutrophils, and liquefaction necrosis are uncommon (187). Splendore in 1908 described a radiate eosinophilic substance in human tissues from patients with sporotrichosis, and Hoeppli in 1932 reported an eosinophilic material around schistosome larvae (126). The Splendore-Hoeppli reaction is indica-

Definitive sporotrichosis diagnosis is based on the isolation and identification of the etiological agent in culture (126). Isolation of S. schenckii is easily obtained after spreading of the clinical specimens on Sabouraud agar with chloramphenicol and on media with cycloheximide, such as mycobiotic agar. After 5 to 7 days of incubation at 25°C, filamentous hyaline colonies start to grow, and after some time, they may develop a dark color, usually in the centers of the colonies (170). To identify an isolate as S. schenckii, one must demonstrate that it undergoes dimorphism by subculturing the fungus on enriched media such as brain heart infusion agar, chocolate agar, and blood agar at 35 to 37°C for 5 to 7 days. Occasional isolates can be difficult to convert and may require multiple subcultures and extended incubation (216). After S. schenckii conversion to the yeast phase, colonies acquire a creamy aspect and a yellow to tan color (170). Environmental Sporothrix strains may also form the yeast phase when grown on appropriate media at 37°C. For this reason, observation of dematiaceous conidia in colonies maintained at 25°C is mandatory (54, 60, 216). For this purpose, slide culture preparations with potato dextrose agar or cornmeal agar are ideal to study S. schenckii conidiogenesis (54). Positive cultures provide the strongest evidence for sporotrichosis, allowing diagnosis of almost all cases of cutaneous disease. Nevertheless, culture diagnosis has significant limitations, mainly in some manifestations of the disease such as S. schenckii induced arthritis, where the collection of material for culture is difficult. Molecular Detection Nonculture methods have been developed to improve the rate and speed of mycological diagnosis (194, 241). Molecular detection of S. schenckii is useful for a rapid diagnosis of sporotrichosis and also valuable in cases of negative cultures due to low fungal burden or secondary infections. Up to now, there has been a scarcity of molecular methods for the detection of S. schenckii DNA from clinical specimens. Sandhu and collaborators (211) reported the development of 21 specific nucleotide probes targeting the large-subunit rRNA genes from several fungi, including S. schenckii. The authors adopted a protocol for DNA extraction from clinical specimens that consists of boiling the specimens in an alkaline

642

BARROS ET AL.

guanidine-phenol-Tris reagent, followed by amplification of a variable region of the 28S rRNA gene with universal primers and amplicon identification using the specific probes. The results displayed a high level of specificity for this test. Some methodologies to identify S. schenckii colonies from pure cultures have been described. Specific probes for fungi with yeast-like morphology in vivo, including all dimorphic fungal pathogens, were developed for the detection of PCR amplicons in an enzyme immunoassay format. S. schenckii DNA was able to hybridize to the probe to detect all dimorphic fungi as well as to its specific probe (137). Specific oligonucleotide primers based on the chitin synthase gene were also developed. This primer was able to detect 10 pg of genomic S. schenckii DNA (110). Primers to distinguish S. schenckii from related species such as Ceratocystis stenoceras, based on the DNA topoisomerase II genes, permitted the amplification of fragments of 663 to 817 bp from S. schenckii and a 660-bp fragment from S. schenckii var. lurei. Another set of primers allowed the amplification of a specific 305-bp fragment from S. schenckii var. lurei (109). These detection systems may be useful as diagnostic tools for the detection of human and animal sporotrichosis. In fact, a PCR assay based on the internal transcriber space in the rRNA gene has been used for the identification of an S. schenckii strain from an atypical case of sporotrichosis (75). Sporotrichin Skin Test The cutaneous sporotrichin skin test detects delayed hypersensitivity, i.e., the cellular immune response, and can be a useful diagnostic tool, but its major usefulness is in epidemiological investigations. This reaction is usually positive in about 90% of confirmed sporotrichosis cases but can also indicate previous infection with the fungus (105). The sporotrichin skin test has been successfully applied to confirm the diagnosis of bulbar conjunctival sporotrichosis after the pathological examination revealed yeast-like cells (113). Epidemiological studies usually involve the sporotrichin skin testing of individuals living or working in a determinate area together with attempts to isolate the fungus from the soil in that area. For instance, this test gave 6.25% positivity in a Mexican state where virulent strains of S. schenckii were isolated from soil (210). On the other hand, 13.67% positivity was found among healthy mine workers from Brazil, although the fungus was not isolated from soil samples from the mines investigated (198). Despite the current use of the sporotrichin skin test in several studies throughout the world, the antigen adopted in these tests lacks standardization. Several studies on sporotrichin in Brazil were performed with a 5 McFarland standard suspension of heat-killed yeast cells (129). A retrospective 10-year study in Mexico used extracted mycelial antigens at a 1:2,000 dilution (28), while another Mexican study diluted yeast-phase antigens at 1:4,000 (210). These variations in antigen production may lead to differences in results. Antibody Detection Several methodologies have been described for the immunological diagnosis of sporotrichosis based on antibody detec-

CLIN. MICROBIOL. REV.

tion in sera from infected patients. Precipitation and agglutination techniques were first adopted. Double immunodiffusion for sporotrichosis does not usually show cross-reactions with sera from patients with chromoblastomycosis or leishmaniasis, infectious diseases with similar clinical manifestations. Immunoelectrophoresis has also been employed, with an anodic arc, called S arc, being observed in all positive cases (2). Tube agglutination and latex agglutination have been utilized for sporotrichosis serodiagnosis since the 1970s, and very good sensitivity (96% and 94%, respectively) and specificity (98 and 100%, respectively) have been observed (26, 41, 112). These tests, however, lack sensitivity in cases of cutaneous sporotrichosis (2, 196) and do not permit the determination of the immunoglobulin isotype involved in the response. Immunoenzymatic assays are currently being used more frequently for serodiagnosis purposes. The publication of an immunoblot assay for serodiagnosis of sporotrichosis dates back to 1989, when molecules of 40 and 70 kDa from exoantigen preparations from the S. schenckii yeast form showed 100% sensitivity and 95% specificity (229). Our group, however, in an attempt to reproduce these data, found high cross-reactivity of this antigenic preparation with sera from patients with paracoccidioidomycosis (unpublished results), which is not an endemic disease in the United States, where the first study was conducted. Antibodies against a concanavalin A binding peptide-rhamnomannan from the S. schenckii yeast cell wall could be detected in an enzyme-linked immunosorbent assay (ELISA), showing 100% sensitivity when 35 serum samples from patients with culture-proven sporotrichosis were tested. However, sera from patients with cutaneous leishmaniasis showed cross-reactions in this assay format (140). This antigenic preparation was further evaluated with sera from 92 patients with sporotrichosis in Rio de Janeiro and 77 heterologous sera, with 90% sensitivity and 80% specificity (23). Other studies showed that the use of different strains in the preparation of the antigen may lead to different sensitivity and specificity results, despite the process of purification of the antigen involved in this methodology. This difference is due to O-glycan residues linked to the molecules. The strain that had better results was the 1099-18 strain obtained from the Mycology Section, Department of Dermatology, Columbia University, New York, NY (24). Efforts to contribute to this field culminated in the development of an enzyme immunoassay with exoantigens produced by a mycelial-phase S. schenckii strain isolated during the Rio de Janeiro epidemic of sporotrichosis (7). This antigen was described by Mendoza and collaborators (161) and had no cross-reaction with antigens and serum samples from patients with coccidioidomycosis, histoplasmosis, or paracoccidioidomycosis. The same antigen was previously used in immunodiffusion and immunoelectrophoresis techniques, without cross-reactions with sera from patients with leishmaniasis or chromoblastomycosis (2). The methodology for production of this antigen is simple and does not require chromatographic steps, making it easy for laboratories with limited resources (7), although more variations can occur with this kind of preparation than with those involving purification procedures. Detection of IgG antibodies against these exoantigens distinguished 90 sera from patients with different clinical forms of

VOL. 24, 2011

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

sporotrichosis, 72 sera from patients with other infectious diseases, and 76 sera from healthy controls, yielding 97% sensitivity and 89% specificity (7). These values are slightly higher than those for the concanavalin A binding fraction of the S. schenckii yeast cell wall (23), and similar observations were made with this purified antigen and crude exoantigens in the serodiagnosis of feline sporotrichosis. Purified antigens exhibited 90% sensitivity and 96% specificity, whereas crude exoantigens showed 96% sensitivity and 98% specificity (65). It has also been demonstrated that in order to improve the overall efficiency of antibody detection for diagnosis and follow-up of human sporotrichosis with the mycelial exoantigen preparation, a search for IgG and IgA antibodies for diagnosis and a search for IgG and IgM antibodies for follow-up purposes must be undertaken (6). Importantly, the results from all antibody detection tests provide a presumptive diagnosis of sporotrichosis and require clinical and epidemiological correlation for an accurate evaluation and determination of the final diagnosis. EPIDEMIOLOGY The natural history of sporotrichosis has been changing gradually in frequency, mode of transmission, and demographic and geographic distributions. It is possible that environmental factors, increased urbanization, and improved diagnostics partly explain the alterations in the profile of the disease. Furthermore, since sporotrichosis is not a reportable disease in most countries, there is little information on the incidence, and the known data are those generated by case publications. Geographic Distribution At the beginning of the last century, sporotrichosis was a common disease in France; it declined after 2 decades and today sporadically appears in Europe. In 2009, an autochthonous case was reported in France (146), and in 2008, another one was reported in the southern region of Italy (49), but these reports are rare and isolated. Despite having been described on five continents, sporotrichosis has a higher prevalence in tropical and temperate zones. The main areas of endemicity are located in Japan (248), India (156), Mexico (144), Brazil (17, 218, 222), Uruguay (45, 47), and Peru (124). In the United States, especially in the Mississippi Valley, outbreaks related to pine seedlings and manipulation of moss have been cited (185). A bibliography search performed in PubMed with the term “Sporothrix schenckii” or “sporotrichosis” for the last 10 years (from January 2001 to November 2010) yielded 407 results. Of these, 83 were excluded (because they were book chapters, reports of other diseases, etc.). Of the remaining 324 references retrieved, 142 were case reports and case series for humans, 22 were case reports and case series for animals, 119 were laboratory studies (molecular studies, antifungal susceptibility tests, environmental studies, etc.), and 41 were various (reviews, letters, comments, or guidelines). These 324 publications were produced by authors from 27 countries. Obviously, these data reflect solely the scientific papers published by workers in these countries, and it is not possible to correlate with disease prevalence (Fig. 6). In the United States, where the incidence of cases is low and usually related to handling

643

sphagnum moss (92), the large number of publications reflects the development of research in the country. The lack of infrastructure for laboratory diagnosis and the lack of research in many countries do not allow for accurate knowledge of disease distribution and environmental sources of the fungus. Newton et al. (175), reporting a case of cutaneous-lymphatic sporotrichosis in a patient from Laos, highlighted the difficulties in laboratory diagnosis of fungal infections in Southeast Asian countries, resulting in a shortage of reports. These authors point out that in regions such as the highlands of China, Laos, Vietnam, and Burma, which have favorable conditions for Sporothrix growth, the prevalence of cases must be much higher than is estimated according to the literature. Distribution by Age and Gender Sporotrichosis affects both genders and all ages. In most regions, the difference in case distribution by age and sex is related to occupation and exposure to the fungus. Among the cases studied by Takenaka et al. (248), 70% were older than 50 years, with an equal division between sexes. In Japan, where that study was conducted, working in agricultural activities has been associated with a higher risk for acquiring sporotrichosis. In Colombia (205) and the southern region of Brazil (10), there was a higher prevalence in men over 40 who were involved in agricultural as well as other high-risk activities. Also in Brazil, in the state of Rio de Janeiro, there was a predominance in women over 40 who were involved in housework and the care of cats with sporotrichosis, the highest risk group for acquiring the disease (18, 218). In Abancay, a rural area of Peru, of the 238 cases studied from 1995 to 1997, 60% occurred in children younger than 15 years old. In that study, no association between age, sex, and exposure to the fungus could be found (184). Transmission and Sources of Infection Sporotrichosis has been traditionally known as “gardeners’ disease,” especially affecting those involved in the cultivation of roses (62). Although in most cases the infection results from inoculation of the fungus by thorn or other pricks, scratches, and other small injuries, a history of trauma can be absent (17, 184, 191). Certain occupational and leisure activities, such as floriculture, horticulture, gardening, fishing, hunting, farming, mining, and others that facilitate exposure to the fungus, have over the years been associated with the transmission of the disease (196). In Uruguay, and more recently in southern Brazil, the hunting of armadillos has been related to cases of sporotrichosis (10, 47). There have been reports of the mycosis following bites or scratches by animals such as cats and squirrels, insect bites, and other injuries. These conditions may either result in direct inoculation or facilitate the entry of the fungus (15, 166). In some situations, such as in an area of endemicity in Peru, the mode of transmission had not been made clear (115, 184) until another study in the same region identified the ownership of cats, outdoor activities, and low socioeconomic status as risk factors for acquiring sporotrichosis (142). Some cases have been reported in laboratory professionals who were infected by manipulating cultures of S. schenckii (48). Interhuman transmission is rare (215). Sporo-

644

BARROS ET AL.

CLIN. MICROBIOL. REV.

FIG. 6. Geographic distribution, by country, of scientific production on sporotrichosis in the 21st century according to the type of publication.

trichosis usually occurs in isolated cases or small outbreaks in families and professionals engaged in high-risk activities. Epidemics are rare and, when they occur, are commonly related to a single source of infection (30). The largest outbreak occurred in Witwatersrand, South Africa, and its description contributed significantly to the current knowledge of sporotrichosis, representing so far the most complete epidemiological investigation of the disease. Between 1941 and 1944, more than 3,000 gold miners were infected by the fungus, which was present in the timber of these mines (96). In the United States, the largest epidemic took place in 1988 and involved a total of 84 cases in 15 states, affecting workers who participated in reforestation programs. The cases were associated with exposure to sphag-

num moss used for the packing of seedlings from a nursery in Pennsylvania (42). Zoonotic Transmission Sporotrichosis has been sporadically associated with scratches or bites from animals such as mice, armadillos, squirrels, dogs, and cats (114). The role of felines in the transmission of the mycosis has gained attention since the 1980s, when Read and Sperling (191) reported an outbreak involving five people exposed to a cat with sporotrichosis. Since then, successive reports from different geographical regions have characterized a new risk group for acquisition of sporotrichosis, composed of

VOL. 24, 2011

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

cat owners and veterinarians (98, 269). The first epidemic of zoonotic sporotrichosis was detected in Rio de Janeiro (17, 222). From 1998 to December 2009, more than 2,000 cases in humans and over 3,000 cases in cats were diagnosed at Instituto de Pesquisa Clínica Evandro Chagas (IPEC)/Fiocruz, representing the largest epidemic of zoonotic transmission of this mycosis ever recorded (19). A study of 178 human cases assessed from 1998 to 2001 showed that of 156 patients who reported professional or household contact with cats with sporotrichosis, 97 had been scratched or bitten by these animals (17). Some authors believe that cats are the only animals with zoonotic transmission potential because of the large amount of yeast cells in the lesions (225, 245). Although dogs have also been affected, they do not seem to have significant zoonotic potential (223). Several studies have been conducted in an attempt to understand why sporotrichosis has reached epidemic proportions in Rio de Janeiro as well as the reason for the high susceptibility of cats to infection by Sporothrix (192, 224–226). However, the lack of studies on environmental, molecular, and feline immune responses against the fungus leave many questions unanswered. Gutierrez-Galhardo et al. (90) investigated the phenotypes and genotypes of Sporothrix schenckii isolates recovered from different clinical forms of the disease. A total of 88 isolates recovered from 59 human cases associated with the epidemic and 29 controls (cases from other Brazilian regions and Spain) were studied. Fingerprinting analysis showed that the Rio de Janeiro epidemic strains were genetically related. Although nine subtypes were found, they were not associated with specific clinical forms. Similar results were obtained by ribosomal DNA sequencing of the internal transcribed spacer sequence. These data suggest that the strains isolated from the cases of sporotrichosis in Rio de Janeiro all originated from a common source. PATHOGENESIS AND CLINICAL FORMS Human Disease Clinical presentations of sporotrichosis may vary according to the immunological status of the host, the load and depth of the inoculum, and the pathogenicity and thermal tolerance of the strain, among other factors (14). Lavalle and Mariat (132) considered that the localized cutaneous form occurs by reinfection of patients who had previously developed immunity to S. schenckii, while the lymphocutaneous form manifests in patients without prior contact with the fungus. According to Rippon (196), continuous exposure to small amounts of conidia in an area of endemicity could gradually confer immunity. An experimental murine model adopted to study the genotypes, virulence, and clinical forms of S. schenckii showed a close relationship between genotype and clinical form. Mice inoculated with isolates from disseminated sporotrichosis presented a shorter time to the onset of the disease and more severe lesions than those inoculated with isolates from lymphocutaneous sporotrichosis. Those inoculated with isolates from the fixed form showed less severe lesions (123). Other studies have reported similar results. Brito et al. (29) studied the virulence of two strains of Sporothrix schenckii, isolated from patients with lymphocutaneous (group 1) or disseminated (group 2) sporotrichosis, by inoculating BALB/c mice. Com-

645

parison of the two groups revealed more severe disease in group 2 mice. The histopathology and large number of viable microorganisms isolated from the spleen confirmed the higher invasive ability of this strain. Furthermore, a decrease of an in vitro specific lymphoproliferative response and IFN-␥ production was observed over time in group 2. These results reinforce the existence of different virulence profiles in S. schenckii strains. According to the location of the lesions, sporotrichosis can be classified into cutaneous, mucosal, and extracutaneous forms. Cutaneous forms. In cutaneous forms, the infection usually appears after minor trauma with disruption of epidermis integrity. After penetrating through the skin, the fungus converts into the yeast form and may remain localized in the subcutaneous tissue or extend along the adjacent lymphatic vessels, constituting the fixed or the lymphocutaneous form, respectively. More rarely it may spread by the hematogenous route, characterizing the disseminated cutaneous form (239). In cases of cat-transmitted sporotrichosis, infection by the yeast form can also occur. This hypothesis is based on the large amount of this form in the lesions from cats associated with transmission without a history of scratches or bites (18). Moreover, the isolation of yeast forms in cats’ claws also favors this idea (226). A molecular study revealed that DNA fingerprints of S. schenckii isolated from the nails and the oral cavities of cats were identical to those of related human samples, suggesting that there is a common infection source for animals and humans in the current epidemic in Rio de Janeiro (192). It is clear that cats act as a vehicle for S. schenckii dissemination. However, the lack of environmental studies on this epidemic and the deficit of information on natural sources that can serve as a reservoir of the fungus do not allow a conclusion to be drawn. The fixed form is represented by a single lesion or a few lesions at the inoculation site, which is often ulcerated with erythematous edges. The morphology can also be vegetative, verrucous, plaque infiltrated, or tuberous, without lymphatic involvement. Some cases may spontaneously regress (4). Most authors report the fixed cutaneous form as the main clinical presentation in children (252). However, the lymphocutaneous form is the most frequent overall, being present in more than 75% of cases (28, 190). The primary lesion is usually located on the extremities, especially hands and forearms, corresponding to the sites most exposed to trauma. Initially, a papule or pustule is followed by formation of a subcutaneous nodule. This lesion, exerting pressure beneath the skin, causes ischemia under the epidermis, evolves into gum, ulcerates, and oozes a purulent secretion. With the progression, secondary lesions arise along the path of regional lymphatics, featuring “sporotrichoid aspect” of the infection. Lymph node involvement or the presence of systemic symptoms is unusual. A disseminated cutaneous form is characterized by multiple skin lesions at noncontiguous sites without extracutaneous involvement. Lesions of the fixed and lymphocutaneous forms may coexist in the same patient. Until the emergence of zoonotic transmission, this form was rare and was caused by hematogenous spread of the fungus, usually associated with immunosuppression (39, 239). In transmission by cats, several inoculations in different locations may occur, during either treatment or play with animals (20). Although there is a dis-

646

BARROS ET AL.

tinct pathogenesis for each situation, it is difficult to identify whether the clinical presentation is due to dissemination from a single lesion or to multiple inoculations. Mucosal form. Some authors consider the mucosal form to be a variant of the cutaneous form. In the nasal mucosa, the lesions often involve the septum, with drainage of bloody secretions and detachment of crusts. In the conjunctiva, the granulomatous lesion is accompanied by a serous-purulent discharge, redness, and presence or not of lid edema (219). Mucosal forms are frequently accompanied by preauricular and submandibular lymph node enlargement (93, 219). These signs can be a consequence of self-inoculation through hands contaminated with the fungus, hematogenous dissemination, and inhalation of conidia (18, 20, 93). Although rare, conjunctival and nasal mucosal involvement has been diagnosed even in pediatric patients (16, 218). In cats with sporotrichosis, the nose is the most affected region and respiratory signs are common (83, 222). As the owners play with their animals in close contact, transmission is made easier. Extracutaneous form. The extracutaneous forms are rare and difficult to diagnose, although they are more frequent after the onset of AIDS (31, 94, 234, 263). After the skin, bone tissue is the most affected. The osteoarticular form may occur by contiguity or hematogenous spread (120, 170). The lesions may vary from small granulomas to large lytic lesions identical to osteomyelitis (125). One or several joints and bones can be involved, as well as tenosynovitis or bursitis (115, 227). In immunocompetent patients, monoarthritis is more frequent than multiple articular involvement. The rarity of musculoskeletal disease in addition to the scarcity of fungal elements in synovial fluid culture and synovial histopathology often delays the diagnosis (11). According to Howell and Toohey, there were only 51 cases of sporothrical arthritis reported in the English literature up to 1998 (102). Primary pulmonary sporotrichosis, resulting from inhalation of the fungus, is usually associated with chronic obstructive pulmonary disease, alcoholism, chronic use of corticosteroids and, immunosuppressive diseases (189). The clinical presentation is similar to that of tuberculosis, and the diagnosis is often delayed due to the rarity of pulmonary involvement. Radiological patterns include cavitary disease, tracheobronchial lymph nodes enlargement, and nodular lesions (183). Systemic sporotrichosis is extremely rare and always associated with immune system deficiency. De Beurmann and Gougerot (51) had suggested that sporotrichosis could be considered an opportunistic disease. Most of their patients had some comorbidity, and all cases with the extracutaneous form presented impaired health or malnutrition. Reports on meningitis associated with Sporothrix infection are not frequent, and these cases are often associated with immunological impairment, mainly after onset of HIV infection. Diagnosis of this form of chronic meningitis is challenging because of the rarity of demonstration of Sporothrix schenckii in smears of cerebrospinal fluid and the difficulty in isolating the yeast on culture. Thus, any other method that provides early and specific diagnosis, such as antibody detection in cerebrospinal fluid, may be helpful in the diagnosis (23, 228). According to Salaki et al. (209), there has been a marked increase in the number of reported cases of meningitis and brain abscess due to fungi and yeasts. This increase is due in part to better diagnostic techniques and greater awareness of the possibility

CLIN. MICROBIOL. REV.

of fungal invasion of the nervous system, but the increase can also be attributed to a growing pool of severely compromised hosts. Besides AIDS, other conditions such as diabetes, alcoholism, granulomatous diseases, cirrhosis, renal transplantation, malignancies, corticosteroids, and use of immunosuppressive agents are commonly reported in patients with extracutaneous sporotrichosis (72, 82, 85). HIV-infected patients with preserved immunity seem to respond to infection by Sporothrix schenckii in the same way as individuals without coinfection (220). In patients with AIDS, sporotrichosis assumes the role of an opportunistic disease, with severe cases, systemic involvement, and often spread to the meninges (263). Skin lesions may be atypical, with a minimal inflammatory response (114). There are cases of disseminated sporotrichosis described in the literature as the first manifestation of AIDS (8) and associated with immune reconstitution inflammatory syndrome (76, 87). Associated signs and symptoms. Erythema nodosum and erythema multiforme have been reported in cases of sporotrichosis by zoonotic transmission. These conditions appear to be associated with a hypersensitivity reaction, resulting from continuous exposure to large amounts of fungus and subclinical reinfections (86, 89). Differential diagnosis. The differential diagnoses should be considered in accordance with the diversity of clinical forms and the morphology of the lesions. In the fixed form, the main differential diagnosis is cutaneous leishmaniasis. A study with application of the Montenegro skin test in 107 cases of sporotrichosis in Rio de Janeiro from 1998 to 2001 produced 48.6% positivity (21). In such cases, only mycological and parasitological examinations can establish a definitive diagnosis. Apart from cutaneous leishmaniasis, other causes, including noninfectious skin ulcers, should be considered. In the lymphocutaneous form, other disorders that present nodular lymphangitis should be investigated (58, 253), particularly mycobacteriosis. Among the mycobacteria, Mycobacterium marinum infection often accompanies lymphocutaneous lesions. There has been a report of sporotrichoid infection caused by Mycobacterium fortuitum in a pregnant woman (208). Nocardiosis (caused mainly by Nocardia brasiliensis), chromoblastomycosis, cryptococcosis, blastomycosis, and cat scratch disease also can be differential diagnoses. Sporotrichosis can also mimic cutaneous bacterial infections, sarcoidosis, lupus vulgaris, tuberculosis, and scrofuloderma, among others (11, 188, 268). These conditions should be differentiated by history, areas of endemicity, and lab tests. Feline Disease De Beurmann et al. (52) experimentally demonstrated the susceptibility of cats to S. schenckii in 1909. However, naturally acquired feline sporotrichosis was reported only in 1952 by Singer and Muncie (236). Sporotrichosis in cats has been considered to be sporadic and transmission to humans to be accidental (61). In 1998 the first epizootic of cat sporotrichosis was detected. By December 2010, over 3,000 cases in cats were diagnosed at Fiocruz/Rio de Janeiro (19). A study by Schubach et al. evaluated 337 cats with sporotrichosis diagnosed by isolation of S. schenckii in culture, 10 asymptomatic cat carriers, and 91 asymptomatic apparently healthy cats in the period

VOL. 24, 2011

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

from 1998 to 2001 (221, 222). In those 4 years of monitoring the epidemic, there was a broad spectrum in clinical presentation, ranging from subclinical infection to single lesions with spontaneous regression to fatal systemic forms. The lymphocutaneous form was determined in only 19.3% of the cases, the involvement of the mucous membranes of the respiratory tract and upper digestive tract in 34.9%, and multiple cutaneous lesions in 39.5% (222). Systemic disease was demonstrated by in vivo detection of S. schenckii in the skin and various internal organs in 10 cats necropsied (225). In cats, unlike in humans, the low frequency of granuloma (12%) and the richness of fungal elements found in the histopathology of the skin demonstrate the increased susceptibility of animals to S. schenckii (222). An evaluation of 142 cats with sporotrichosis regarding feline immunodeficiency virus (FIV) and/or feline leukemia virus (FeLV) infection revealed 21.8% positivity (222). There were no significant differences in clinical and laboratory profiles between animals coinfected and not coinfected with FIV and/or FeLV. Unlike in humans, in whom disseminated sporotrichosis typically affects immunocompromised individuals, systemic disease in cats was frequent and was not associated with immunodeficiency caused by FIV and/or FeLV. TREATMENT Potassium iodide has been traditionally used in the treatment of sporotrichosis since the early 20th century, with satisfactory results (51). Some suggest that this salt acts on the resolution of granulomas through increased proteolysis (196), while others claim that it promotes an increase of phagocytosis. However, the exact mechanism of action remains unknown. Due to adverse effects related to this medication, in the 1990s the azole compounds were introduced, and itraconazole is currently the first-choice treatment (115). Itraconazole has been used effectively and safely in most cases of sporotrichosis, with low toxicity and good tolerance, even in long-term treatments. The dose varies from 100 to 200 mg/day orally for cutaneous and lymphocutaneous forms. A dose of 400 mg/day orally should be administered for cutaneous forms with a poor initial response to a lower dosage, for osteoarticular forms, for pulmonary forms, and as suppressive therapy in immunocompromised patients after induction with amphotericin B. For children weighing up to 20 kg, 5 to 10 mg/kg/day is recommended. Itraconazole is metabolized by the cytochrome P450 3A4 and has numerous drug interactions associated with the inhibition or induction of this system. The drug is contraindicated in pregnancy, and women of childbearing age should be counseled in favor of effective contraceptive methods. The safety and efficacy of itraconazole were evaluated in 645 patients with culture-proven sporotrichosis from Rio de Janeiro, Brazil. Six hundred ten (94.6%) of the patients were cured with itraconazole (50 to 400 mg/day), 547 with 100 mg/day, 59 with 200 to 400 mg/day, and 4 children with 50 mg/day. Four hundred sixty-two (71.6%) completed clinical follow-up for 3 to 6 months, and all remained cured. This study highlighted the good response to a minimal dose of itraconazole (55). Several reports have demonstrated the excellent efficacy and tolerability of the drug, even for the disseminated cutaneous and extracutaneous forms (119, 149, 165, 239, 268). The intermittent

647

administration of itraconazole in pulses of 400 mg/day orally for 1 week has also been studied (27). A randomized controlled study comparing the safety and efficacy of an itraconazole pulse regimen to a continuous regimen for cutaneous sporotrichosis found similar results for both groups (238). Potassium iodide and itraconazole are contraindicated in pregnant women. These patients can use thermotherapy with daily application of local heat (42 to 43°C) through a hot water bag, a source of infrared, or a similar method until healing of the lesions (115). The mechanism of action of local heat has been demonstrated in the laboratory. When cells of S. schenckii in serum are incubated with neutrophils at 40°C and 37°C, there is no difference related to phagocytosis in the two groups. However, once the cells are phagocytosed, the death rate of the fungus is higher at 40°C than at 37°C (99). Amphotericin B has been given for initial treatment of disseminated forms, particularly in immunocompromised subjects (115). Patients coinfected with HIV should subsequently receive suppressive therapy with itraconazole until immune system restoration. In pregnant women, amphotericin B may be used after 12 weeks of pregnancy, but this medication is to be reserved for pulmonary and disseminated forms for which treatment cannot be delayed. Fluconazole is less effective than itraconazole and is given to patients who do not tolerate or have drug interactions with itraconazole. Ketoconazole, in addition to having greater toxicity, has not demonstrated a good response (115). Susceptibility studies concerning good in vitro activity of terbinafine and posaconazole against Sporothrix schenckii have been encouraging (157, 235). The three studies using terbinafine in human patients demonstrated a good efficacy for doses ranging from 250 to 1,000 mg/day (43, 73, 74). A study comparing 250 mg/day of terbinafine to 100 mg/day of itraconazole resulted in healing in 92.7% and 92% of patients, respectively, indicating terbinafine to be an effective and well-tolerated option for the treatment of cutaneous sporotrichosis (73). However, the cost, which is even higher than that of itraconazole, is a barrier for developing countries. Posaconazole is a broadspectrum triazole, but its use in clinical sporotrichosis is yet to be evaluated (214, 235). Susceptibility testing with voriconazole, ravuconazole, micafungin, and other new antifungal agents is still incipient, and so far there is no indication for use of these drugs for the treatment of sporotrichosis (9, 169, 178, 233). There are studies showing different results for the same antifungal agent according to geographic area, the species of Sporothrix, and the method used (9, 88, 169). Antifungal Susceptibility Tests Nowadays, there is an ever-increasing interest in testing the susceptibilities of filamentous and yeast fungal pathogens to the available antifungal drugs (80). Of the available methods, two are standardized for S. schenckii testing. The Clinical and Laboratory Standards Institute (CLSI) proposes a microdilution test with a test inoculum of 0.4 ⫻ 104 to 5 ⫻ 104 CFU/ml, rendered from a 7-day-old S. schenckii filamentous colony, used to inoculate RPMI 1640 medium, where antifungal drugs are serially diluted and, after 46 to 50 h of incubation at 35°C without agitation, MICs determined (46). Another protocol for S. schenckii antifungal susceptibility testing was elaborated by the European Committee for Antimicrobial Susceptibility

648

BARROS ET AL.

CLIN. MICROBIOL. REV.

TABLE 1. MIC ranges for 13 antifungal drugs against S. schenckii described in various publicationsa Reference

64 106 155 172 121 179 169 81 258 122 10 157 91 152 208 235

Location

USA UK Not reported Japan Brazil USA Peru Mexico Brazil Brazil Peru Brazil Brazil/Spain Several Not reported Several

MIC (␮g/ml)b

No. of strains analyzed

AMB

FLC

ITR

KET

VOR

5 10 100 7 30 3 22 15 34 43 19 12 88 92 10 62

0.5–16 0.5–4 0.25–2 1–2 NT 1–4 NT NT 1–4 0.5–8 0.06–⬎16 NT 0.12–4 0.5–32 NT 0.03–16

64 NT NT ⬎64 NT NT NT ⬎64 ⬎64 NT ⬎64 NT 64 128 NT NT

0.03–16 0.06–⬎16 0.03–8 0.5–1 0.06–4 NT 0.06–1 0.25–2 0.25–⬎16 0.25–4 0.25–1 0.2–1.75 0.12–⬎8 0.5–32 0.5–2 0.03–2

0.25–4 NT NT NT NT NT NT NT 0.03–16 NT 0.06–0.5c NT NT 0.06–8 NT NT

NT 0.12–⬎16 0.5–8 NT NT ⬎16 2–16 1–4 0.5–⬎16 NT 4–16 NT 1–⬎8 0.5–32 NT NT

a

Results were obtained by microdilution reference methods unless otherwise specified. AMB, amphotericin B; FLC, fluconazole; ITR, itraconazole; KET, ketoconazole; VOR, voriconazole; MFG, micafungin; ADF, anidulafungin; TRB, terbinafine; 5FC, 5-fluorocytosine; RVC, ravuconazole; POS, posaconazole; ABC, albaconazole; EBC, eberconazole; NT, not tested. c The MIC was determined by the Etest method. b

Testing (EUCAST), which uses final test inoculums of 1 ⫻ 105 to 2.5 ⫻ 105 CFU/ml obtained from 2- to 5-day-old cultures to inoculate microplates with RPMI 1640 supplemented with 2% glucose. The endpoint is read visually after a 48 h of incubation at 35 ⫾ 2°C (201). Commercial methods for S. schenckii susceptibility testing, however, have been adopted by some investigators. The Etest technique shows average agreement rates of 77.5% and 87.8% for amphotericin B and fluconazole, respectively, but for itraconazole and voriconazole agreement is lower, with values of 56.4% and 54.5%, respectively. MIC values for itraconazole and voriconazole determined by Etest were significantly lower than those obtained by microdilution (87). The Sensititre YeastOne and ATB Fungus 2 methods compared with CLSI M38-A2 show high agreement with microdilution for fluconazole, itraconazole, and 5-fluorocytosine, but for amphotericin B and voriconazole, agreement is lower (9). Up to now, breakpoints have not been established for molds, including the dimorphic fungus S. schenckii. However, studies showed that MICs of below 1 ␮g/ml are usually found for Scedosporium apiospermum and Paecilomyces lilacinus with posaconazole and voriconazole, for Alternaria spp. and Bipolaris spicifera with posaconazole, voriconazole, and itraconazole, for zygomycetes with amphotericin B and posaconazole, and for Aspergillus spp. with posaconazole, voriconazole, itraconazole, amphotericin, and caspofungin. Based on this, isolates can be grouped as susceptible (MIC ⱕ 1 ␮g/ml), intermediate (MIC ⫽ 2 ␮g/ml), or resistant (MIC ⱖ 4 ␮g/ml) only for analytical purposes. Breakpoints with proven relevance have yet to be identified by the regulatory agencies (46). Despite the use of the nonparasitic S. schenckii mycelial phase in standardized methods, some studies have compared susceptibilities for the mycelial and yeast fungal morphological phases. Agreements of above 80% were found for amphotericin B, fluconazole, itraconazole, ravuconazole, and terbinafine. For posaconazole, an agreement of 76.5% was found for mycelial and yeast forms, and only 47.6% was verified for

voriconazole (87) in a study with Brazilian and Spanish S. schenckii strains. Another study found similar MICs for itraconazole and terbinafine, although the MIC of amphotericin B was slightly higher for mycelia (122). It has been also demonstrated that micafungin is more active against the mycelial than the yeast S. schenckii form (172). Table 1 displays the MIC range for several drugs tested in 16 different studies concerning S. schenckii susceptibility profiles. Since the methodologies used in these studies may have slight differences, comparison of results is difficult. However, some conclusions can be reached. MICs for S. schenckii show high variability (63), as we can see in studies on itraconazole susceptibility. S. schenckii susceptibility to this triazole drug ranges from 0.03 to more than 16 ␮g/ml. Although some strains can show resistance to this drug, geometric means commonly are low, ranging from 0.4 ␮g/ml (9) to 4.08 ␮g/ml (257). In general, despite the use of fluconazole as a therapeutic agent in some human and veterinary sporotrichosis cases (31, 50), this antifungal does not inhibit S. schenckii growth in vitro (9, 63, 90, 152, 172, 257). Some S. schenckii strains are susceptible to voriconazole in vitro, since MIC values of as low as 0.12 ␮g/ml have been found (107); however, the high geometric mean MIC values, ranging from 6.50 ␮g/ml (155) to 13.2 ␮g/ml (152), suggest that most isolates are resistant in vitro to this drug. High micafungin MIC values have been reported by all but one study (152, 172, 207, 257). However, the interaction of micafungin and itraconazole shows a synergy against S. schenckii (207). The geographic distribution of S. schenckii strains also appears to play an important role in antifungal susceptibility. Strains from the sporotrichosis area of endemicity in Rio de Janeiro are more susceptible to itraconazole and terbinafine than strains from Spain or Sa˜o Paulo, a neighbor state Rio de Janeiro in Brazil (90). Low terbinafine MIC values were described for strains from Venezuela, and high minimal fungicidal concentration values for posaconazole were found in Peruvian strains (235). This behavior must be related to differences in susceptibility of the newly characterized species

VOL. 24, 2011

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

649

TABLE 1—Continued MIC (␮g/ml)b MFG

ADF

TRB

5FC

RVC

POS

ABC

EBC

NT NT NT 0.5–1 NT NT NT NT ⬎128 NT NT NT NT 256 2–64 NT

NT NT NT NT NT 2–4 NT NT NT NT NT NT NT NT NT NT

NT NT NT NT 0.007–0.5 NT NT NT 0.12–4 0.015–1 NT 0.05–0.875 0.02–⬎16 0.06–4 NT 0.03–1

NT NT NT NT NT NT NT NT 16–⬎64 NT 4–⬎64 NT 4–⬎64 1–128 NT NT

NT NT NT NT NT NT NT 0.5–4 1–16 NT NT NT 0.12–⬎8 0.06–32 NT NT

NT NT NT NT NT NT NT 1–4 NT NT NT NT 0.06–⬎8 0.25–16 NT 0.03–2

NT NT NT NT NT NT NT NT 0.12–16 NT NT NT NT 0.25–32 NT NT

NT NT NT NT NT NT NT NT 0.06–4 NT NT NT NT 0.06–32 NT NT

from the Sporothrix complex. In fact, S. mexicana shows high MIC values of amphotericin B and several azoles, being susceptible to terbinafine only. S. brasiliensis also is more susceptible to azole drugs than the other species (152). PREVENTION Most cases of sporotrichosis occur when the fungus is introduced through a cut or puncture in the skin while handling vegetation or organic matter containing the fungal spores. Control measures include wearing gloves and long sleeves during high-risk activities such as handling sphagnum moss, wires, rose bushes, hay bales, conifer (pine) seedlings, or other materials that may facilitate the exposure to the fungus. A study by Hajjeh et al. (92) showed that the risk of sporotrichosis increased significantly with the duration of working with sphagnum moss, in particular with filling topiaries, and with having less gardening experience. Wearing gloves was protective. It is also advisable to wear heavy boots to prevent puncture wounds. Sporotrichosis in cats requires preventive measures to avoid transmission within the species and from animals to humans. Due to the itinerant nature of cats, where males frequently engage in disputes over females, infection is quite common. Cats with sporotrichosis should be correctly treated and kept isolated in a proper place. Any physical contact with the animal should be avoided until complete healing of the lesions. When handling the sick cat, during either injury treatment or medication administration, protocols must be adopted to reduce exposure to the fungus, such as using latex gloves. Another important measure is not to abandon the animal, as this facilitates the dissemination of the disease. In the case of cats with extensive lesions and no possibility of treatment, euthanasia and cremation of the body should be standard procedures in a veterinary health centers. Castration encumbers the instinct for hunting, mating, and roaming the neighborhood, therefore reducing the chance of transmission of the mycosis. Some intervention in the environment may be necessary, such as cleaning yards and removing remnants of construction materials and decaying organic matter debris. Only the treatment of

sick cats and measures regarding feline sporotrichosis will afford zoonotic transmission control (19). ACKNOWLEDGMENTS We thank Rosani Santos Reis, who kindly provided Fig. 5. This study was partially supported by the Conselho Nacional de Desenvolvimento Cientı´fico e Technolo ´gico (CNPq), Brazil, by the programa de Apoio a Pesquisa Estrate´gica em Sau ´de (PAPES)— Fundac¸˜ao Oswaldo Cruz (Fiocruz)/CNPq, and by the Fundac¸˜ao de Apoia a Pesquisa do Estado do Rio de Janeiro (FAPERJ), Brazil. REFERENCES 1. Ahearn, D. G., and W. Kaplan. 1969. Occurrence of Sporotrichum schenckii on a cold-stored meat product. Am. J. Epidemiol. 89:116–124. 2. Albornoz, M. B., E. Villanueva, and E. D. Torres. 1984. Application of immunoprecipitation techniques to the diagnosis of cutaneous and extracutaneous forms of sporotrichosis. Mycopathologia 85:177–183. 3. Albuquerque, P. C., et al. 2008. Vesicular transport in Histoplasma capsulatum: an effective mechanism for trans-cell wall transfer of proteins and lipids in ascomycetes. Cell. Microbiol. 10:1695–1710. 4. Almeida, H. L., Jr, C. B. Lettnin, J. L. Barbosa, and M. C. Dias. 2009. Spontaneous resolution of zoonotic sporotrichosis during pregnancy. Rev. Inst. Med. Trop. Sao Paulo 51:237–238. 5. Almeida-Paes, R., et al. 2009. Growth conditions influence melanization of Brazilian clinical Sporothrix schenckii isolates. Microbes Infect. 11:554–562. 6. Almeida-Paes, R., M. A. Pimenta, P. C. Monteiro, J. D. Nosanchuk, and R. M. Zancope-Oliveira. 2007. Immunoglobulins G, M, and A against Sporothrix schenckii exoantigens in patients with sporotrichosis before and during treatment with itraconazole. Clin. Vaccine Immunol. 14:1149–1157. 7. Almeida-Paes, R., et al. 2007. Use of mycelial-phase Sporothrix schenckii exoantigens in an enzyme-linked immunosorbent assay for diagnosis of sporotrichosis by antibody detection. Clin. Vaccine Immunol. 14:244–249. 8. Al-Tawfiq, J. A., and K. K. Wools. 1998. Disseminated sporotrichosis and Sporothrix schenckii fungemia as the initial presentation of human immunodeficiency virus infection. Clin. Infect. Dis. 26:1403–1406. 9. Alvarado-Ramirez, E., and J. M. Torres-Rodriguez. 2007. In vitro susceptibility of Sporothrix schenckii to six antifungal agents determined using three different methods. Antimicrob. Agents Chemother. 51:2420–2423. 10. Alves, S. H., et al. 2010. Sporothrix schenckii associated with armadillo hunting in Southern Brazil: epidemiological and antifungal susceptibility profiles. Rev. Soc. Bras. Med. Trop. 43:523–525. 11. Appenzeller, S., et al. 2006. Sporothrix schenckii infection presented as monoarthritis: report of two cases and review of the literature. Clin. Rheumatol. 25:926–928. 12. Area-Lea ˜o, A. E., and M. Goto. 1946. Esporotricose. Observac¸˜ao e estudo de um novo caso. Hospital 30:409–417. 13. Arenas, R., D. Miller, and P. Campos-Macias. 2007. Epidemiological data and molecular characterization (mtDNA) of Sporothrix schenckii in 13 cases from Mexico. Int. J. Dermatol. 46:177–179. 14. Arrillaga-Moncrieff, I., et al. 2009. Different virulence levels of the species of Sporothrix in a murine model. Clin. Microbiol. Infect. 15:651–655.

650

BARROS ET AL.

15. Barba Borrego, J. A., J. Mayorga, and V. M. Tarango-Martinez. 2009. Esporotricosis linfangitica bilateral y simultanea. Rev. Iberoam. Micol. 26:247–249. 16. Barros, M. B., et al. 2008. Endemic of zoonotic sporotrichosis: profile of cases in children. Pediatr. Infect. Dis. J. 27:246–250. 17. Barros, M. B. L., A. O. Schubach, A. C. do Valle, et al. 2004. Cat-transmitted sporotrichosis epidemic in Rio de Janeiro, Brazil: description of a series of cases. Clin. Infect. Dis. 38:529–535. 18. Barros, M. B. L., A. O. Schubach, T. M. Schubach, B. Wanke, and S. R. Lambert-Passos. 2008. An epidemic of sporotrichosis in Rio de Janeiro, Brazil: epidemiological aspects of a series of cases. Epidemiol. Infect. 136: 1192–1196. 19. Barros, M. B. L., et al. 2010. Esporotricose: a evoluc¸˜ao e os desafios de uma epidemia. Rev. Panam. Salud Publ. 27:455–460. 20. Barros, M. B. L., et al. 2003. Sporotrichosis with widespread cutaneous lesions: report of 24 cases related to transmission by domestic cats in Rio de Janeiro, Brazil. Int. J. Dermatol. 42:677–681. 21. Barros, M. B. L., A. Schubach, A. C. Francesconi-do-Valle, et al. 2005. Positive Montenegro skin test among patients with sporotrichosis in Rio De Janeiro. Acta Trop. 93:41–47. 22. Berbee, M. L., and J. W. Taylor. 1992. Convergence in ascospore discharge mechanism among pyrenomycete fungi based on 18S ribosomal RNA gene sequence. Mol. Phylogenet. Evol. 1:59–71. 23. Bernardes-Engemann, A. R., et al. 2005. Development of an enzyme-linked immunosorbent assay for the serodiagnosis of several clinical forms of sporotrichosis. Med. Mycol. 43:487–493. 24. Bernardes-Engemann, A. R., et al. 2009. A comparative serological study of the SsCBF antigenic fraction isolated from three Sporothrix schenckii strains. Med. Mycol. 47:874–878. 25. Betancourt, S., L. J. Torres-Bauza, and N. Rodriguez-Del Valle. 1985. Molecular and cellular events during the yeast to mycelium transition in Sporothrix schenckii. Sabouraudia 23:207–218. 26. Blumer, S. O., L. Kaufman, W. Kaplan, D. W. McLaughlin, and D. E. Kraft. 1973. Comparative evaluation of five serological methods for the diagnosis of sporotrichosis. Appl. Microbiol. 26:4–8. 27. Bonifaz, A., L. Fierro, A. Saul, and R. M. Ponce. 2008. Cutaneous sporotrichosis. Intermittent treatment (pulses) with itraconazole. Eur. J. Dermatol. 18:61–64. 28. Bonifaz, A., et al. 2007. Sporotrichosis in childhood: clinical and therapeutic experience in 25 patients. Pediatr. Dermatol. 24:369–372. 29. Brito, M. M., et al. 2007. Comparison of virulence of different Sporothrix schenckii clinical isolates using experimental murine model. Med. Mycol. 45:721–729. 30. Bustamante, B., and P. E. Campos. 2001. Endemic sporotrichosis. Curr. Opin. Infect. Dis. 14:145–149. 31. Callens, S. F., et al. 2006. Pulmonary Sporothrix schenckii infection in a HIV positive child. J. Trop. Pediatr. 52:144–146. 32. Carlos, I. Z., M. F. Sassa, D. B. da Graca Sgarbi, M. C. Placeres, and D. C. Maia. 2009. Current research on the immune response to experimental sporotrichosis. Mycopathologia 168:1–10. 33. Carlos, I. Z., D. B. Sgarbi, J. Angluster, C. S. Alviano, and C. L. Silva. 1992. Detection of cellular immunity with the soluble antigen of the fungus Sporothrix schenckii in the systemic form of the disease. Mycopathologia 117:139–144. 34. Carlos, I. Z., D. B. Sgarbi, and M. C. Placeres. 1998. Host organism defense by a peptide-polysaccharide extracted from the fungus Sporothrix schenckii. Mycopathologia 144:9–14. 35. Carlos, I. Z., D. B. Sgarbi, G. C. Santos, and M. C. Placeres. 2003. Sporothrix schenckii lipid inhibits macrophage phagocytosis: involvement of nitric oxide and tumour necrosis factor-alpha. Scand. J. Immunol. 57:214–220. 36. Carlos, I. Z., et al. 1994. Disturbances in the production of interleukin-1 and tumor necrosis factor in disseminated murine sporotrichosis. Mycopathologia 127:189–194. 37. Carmichael, J. W. 1962. Chrysosporium and some other aleuriosporic Hyphomycetes. Can. J. Bot. 40:1137–1173. 38. Carvalho, L. P., et al. 2007. Differential immune regulation of activated T cells between cutaneous and mucosal leishmaniasis as a model for pathogenesis. Parasite Immunol. 29:251–258. 39. Carvalho, M. T., et al. 2002. Disseminated cutaneous sporotrichosis in a patient with AIDS: report of a case. Rev. Soc. Bras. Med. Trop. 35:655–659. 40. Casadevall, A. 2006. Cards of virulence and the global virulome for humans. Microbe 1:359–364. 41. Casserone, S., I. A. Conti-Diaz, E. Zanetta, and M. E. P. Pereira. 1983. Serologia de la esporotricosis cutaˆnea. Sabouraudia 21:317–321. 42. Centers for Disease Control and Prevention. 1988. Multistate outbreak of sporotrichosis in seedling handlers. Morb. Mortal. Wkly. Rep. 37:652–653. 43. Chapman, S. W., et al. 2004. Comparative evaluation of the efficacy and safety of two doses of terbinafine (500 and 1000 mg day(⫺1)) in the treatment of cutaneous or lymphocutaneous sporotrichosis. Mycoses 47: 62–68. 44. Chua, S. S., M. Momany, L. Mendoza, and P. J. Szaniszlo. 1994. Identifi-

CLIN. MICROBIOL. REV.

45.

46.

47. 48. 49.

50.

51. 52. 53. 54.

55. 56.

57.

58. 59.

60.

61.

62. 63.

64.

65.

66.

67.

68. 69.

70.

71. 72. 73.

cation of three chitin synthase genes in the dimorphic fungal pathogen Sporothrix schenckii. Curr. Microbiol. 29:151–156. Civila, E. S., J. Bonasse, I. A. Conti-Diaz, and R. A. Vignale. 2004. Importance of the direct fresh examination in the diagnosis of cutaneous sporotrichosis. Int. J. Dermatol. 43:808–810. Clinical and Laboratory Standards Institute. 2008. Reference method for broth dillution antifungal susceptibility testing of filamentous fungi; approved standard, 2nd ed. Clinical and Laboratory Standards Institute, Wayne, PA. Conti Diaz, I. A. 1989. Epidemiology of sporotrichosis in Latin America. Mycopathologia 108:113–116. Cooper, C. R., D. M. Dixon, and I. F. Salkin. 1992. Laboratory-acquired sporotrichosis. J. Med. Vet. Mycol. 30:169–171. Criseo, G., G. Malara, O. Romeo, and A. Puglisi Guerra. 2008. Lymphocutaneous sporotrichosis in an immunocompetent patient: a case report from extreme southern Italy. Mycopathologia 166:159–162. Crothers, S. L., S. D. White, P. J. Ihrke, and V. K. Affolter. 2009. Sporotrichosis: a retrospective evaluation of 23 cases seen in northern California (1987–2007). Vet. Dermatol. 20:249–259. De Beurmann, L., and H. Gougerot. 1912. Les sporotrichoses. Librarie Fe´lix Alcan, Paris, France. De Beurmann, L., H. Gougerot, and A. B. Vaucher. 1909. Sporotichoses cutane´es du chat. C. R. Soc. Biol. 66:370–372. de Hoog, G. S., and J. Guarro. 1995. Atlas of clinical fungi. Centraalbureau voor Shimmelcultures, Baarn and Delft, The Netherlands. de Hoog, G. S., and R. G. Vitale. 2007. Bipolaris, Exophiala, Scedosporium, Sporothrix and other dematiaceous fungi, p. 1898–1917. In P. R. Murray, E. J. Baron, J. H. Jorgensen, M. L. Landry, and M. A. Pfaller (ed.), Manual of clinical microbiology, 9th ed., vol. 2. ASM Press, Washington, DC. de Lima Barros, M. B., et al. 2011. Treatment of cutaneous sporotrichosis with itraconazole—study of 645 patients. Clin. Infect. Dis. 52:e200–e206. de Meyer, E. M., et al. 2008. Taxonomy and phylogeny of new wood- and soil-inhabiting Sporothrix species in the Ophiostoma stenoceras-Sporothrix schenckii complex. Mycologia 100:647–661. de Miranda, L. H., L. P. Quintella, I. B. dos Santos, et al. 2009. Histopathology of canine sporotrichosis: a morphological study of 86 cases from Rio de Janeiro (2001-2007). Mycopathologia 168:79–87. DiNubile, M. J. 2008. Nodular lymphangitis: a distinctive clinical entity with finite etiologies. Curr. Infect. Dis. Rep. 10:404–410. Dixon, D. M., R. A. Duncan, and N. J. Hurd. 1992. Use of a mouse model to evaluate clinical and environmental isolates of Sporothrix spp. from the largest U.S. epidemic of sporotrichosis. J. Clin. Microbiol. 30:951–954. Dixon, D. M., et al. 1991. Isolation and characterization of Sporothrix schenckii from clinical and environmental sources associated with the largest U.S. epidemic of sporotrichosis. J. Clin. Microbiol. 29:1106–1113. Dunstan, R. W., R. F. Langham, K. A. Reimann, and P. S. Wakenell. 1986. Feline sporotrichosis: a report of five cases with transmission to humans. J. Am. Acad. Dermatol. 15:37–45. Engle, J., J. Desir, and J. M. Bernstein. 2007. A rose by any other name. Skinmed 6:139–141. Espinel-Ingroff, A., et al. 1995. Comparative and collaborative evaluation of standardization of antifungal susceptibility testing for filamentous fungi. Antimicrob. Agents Chemother. 39:314–319. Feeney, K. T., I. H. Arthur, A. J. Whittle, S. A. Altman, and D. J. Speers. 2007. Outbreak of sporotrichosis, Western Australia. Emerg. Infect. Dis. 13:1228–1231. Fernandes, G. F., et al. 2011. Serodiagnosis of sporotrichosis infection in cats by enzyme-linked immunosorbent assay using a specific antigen, SsCBF, and crude exoantigens. Vet. Microbiol. 147:445–449. Fernandes, K. S., A. L. Coelho, L. M. Lopes Bezerra, and C. Barja-Fidalgo. 2000. Virulence of Sporothrix schenckii conidia and yeast cells, and their susceptibility to nitric oxide. Immunology 101:563–569. Fernandes, K. S., H. L. Mathews, and L. M. Lopes Bezerra. 1999. Differences in virulence of Sporothrix schenckii conidia related to culture conditions and cell-wall components. J. Med. Microbiol. 48:195–203. Fernandes, K. S., et al. 2008. Detrimental role of endogenous nitric oxide in host defence against Sporothrix schenckii. Immunology 123:469–479. Figueiredo, C. C., O. C. De Lima, L. De Carvalho, L. M. Lopes-Bezerra, and V. Morandi. 2004. The in vitro interaction of Sporothrix schenckii with human endothelial cells is modulated by cytokines and involves endothelial surface molecules. Microb. Pathog. 36:177–188. Figueiredo, C. C., P. M. Deccache, L. M. Lopes-Bezerra, and V. Morandi. 2007. TGF-beta1 induces transendothelial migration of the pathogenic fungus Sporothrix schenckii by a paracellular route involving extracellular matrix proteins. Microbiology 153:2910–2921. Flournoy, D. J., J. B. Mullins, and R. J. McNeal. 2000. Isolation of fungi from rose bush thorns. J. Okla. State Med. Assoc. 93:271–274. Fonseca-Reyes, S., et al. 2007. Extracutaneous sporotrichosis in a patient with liver cirrhosis. Rev. Iberoam. Micol. 24:41–43. Francesconi, G., A. C. Francesconi do Valle, et al. 2011. Comparative study of 250 mg/day terbinafine and 100 mg/day itraconazole for the treatment of cutaneous sporotrichosis. Mycopathologia 171:349–354.

VOL. 24, 2011 74. Francesconi, G., A. C. Valle, S. Passos, R. Reis, and M. C. Galhardo. 2009. Terbinafine (250 mg/day): an effective and safe treatment of cutaneous sporotrichosis. J. Eur. Acad. Dermatol. Venereol. 23:1273–1276. 75. Fujii, H., et al. 2008. A case of atypical sporotrichosis with multifocal cutaneous ulcers. Clin. Exp. Dermatol. 33:135–138. 76. Galhardo, M. C., et al. 2010. Sporothrix schenckii meningitis in AIDS during immune reconstitution syndrome. J. Neurol. Neurosurg Psych. 81:696–699. 77. Garrison, R. G., K. S. Boyd, and F. Mariat. 1975. Ultrastructural studies of the mycelium-to yeast transformation of Sporothrix schenckii. J. Bacteriol. 124:959–968. 78. Gezuele, E., and D. Da Rosa. 2005. Relevancia del cuerpo asteroide esporotricosico en el diagnostico rapido de la esporotricosis. Rev. Iberoam. Micol. 22:147–150. 79. Ghosh, A., P. K. Maity, B. M. Hemashettar, V. K. Sharma, and A. Chakrabarti. 2002. Physiological characters of Sporothrix schenckii isolates. Mycoses 45:449–454. 80. Gonzalez, G. M., A. W. Fothergill, D. A. Sutton, M. G. Rinaldi, and D. Loebenberg. 2005. In vitro activities of new and established triazoles against opportunistic filamentous and dimorphic fungi. Med. Mycol. 43:281–284. 81. Gordon, M. A., E. W. Lapa, M. S. Fitter, and M. Lindsay. 1980. Susceptibility of zoopathogenic fungi to phytoalexins. Antimicrob. Agents Chemother. 17:120–123. 82. Gottlieb, G. S., C. F. Lesser, K. K. Holmes, and A. Wald. 2003. Disseminated sporotrichosis associated with treatment with immunosuppressants and tumor necrosis factor-alpha antagonists. Clin. Infect. Dis. 37:838–840. 83. Gremiao, I. D., et al. 2009. Intralesional amphotericin B in a cat with refractory localised sporotrichosis. J. Feline Med. Surg. 11:720–723. 84. Guarro, J., J. Gene, and A. M. Stchigel. 1999. Developments in fungal taxonomy. Clin. Microbiol. Rev. 12:454–500. 85. Gullberg, R. M., A. Quintanilla, M. L. Levin, J. Williams, and J. P. Phair. 1987. Sporotrichosis: recurrent cutaneous, articular, and central nervous system infection in a renal transplant recipient. Rev. Infect. Dis. 9:369–375. 86. Gutierrez-Galhardo, M. C., et al. 2005. Erythema multiforme associated with sporotrichosis. J. Eur. Acad. Dermatol. Venereol. 19:507–509. 87. Gutierrez-Galhardo, M. C., A. C. do Valle, et al. 2010. Disseminated sporotrichosis as a manifestation of immune reconstitution inflammatory syndrome. Mycoses 53:78–80. 88. Gutierrez-Galhardo, M. C., R. M. Zancope-Oliveira, A. Monzon, J. L. Rodriguez-Tudela, and M. Cuenca-Estrella. 2010. Antifungal susceptibility profile in vitro of Sporothrix schenckii in two growth phases and by two methods: microdilution and E-test. Mycoses 53:227–231. 89. Gutierrez Galhardo, M. C., et al. 2002. Erythema nodosum associated with sporotrichosis. Int. J. Dermatol. 41:114–116. 90. Gutierrez Galhardo, M. C., et al. 2008. Molecular epidemiology and antifungal susceptibility patterns of Sporothrix schenckii isolates from a cattransmitted epidemic of sporotrichosis in Rio de Janeiro, Brazil. Med. Mycol. 46:141–151. 91. Haddad, V. J., H. A. Miot, L. D. Bartoli, C. Cardoso Ade, and R. M. de Camargo. 2002. Localized lymphatic sporotrichosis after fish-induced injury (Tilapia sp.). Med. Mycol. 40:425–427. 92. Hajjeh, R., et al. 1997. Outbreak of sporotrichosis among tree nursery workers. J. Infect. Dis. 176:499–504. 93. Hampton, D. E., A. Adesina, and J. Chodosh. 2002. Conjunctival sporotrichosis in the absence of antecedent trauma. Cornea 21:831–833. 94. Hardman, S., I. Stephenson, D. R. Jenkins, M. J. Wiselka, and E. M. Johnson. 2005. Disseminated Sporothix schenckii in a patient with AIDS. J. Infect. 51:e73–77. 95. Hektoen, L., and C. F. Perkins. 1900. Refractory subcutaneous abscess caused by Sporothrix schenckii: a new pathogenic fungus. J. Exp. Med. 5:77–91. 96. Helm, M., and C. Berman. 1947. The clinical, therapeutic and epidemiological features of the sporotrichosis infection on the mines, p. 59–67. In Sporotrichosis infection on mines of the Witwatersrand. Proceedings of the Transvaal Mine Medical Officers’ Association, December 1944, Johannesburg, South Africa. The Transvaal Chamber of Mines, Johannesburg, South Africa. 97. Hintz, W. E. 1999. Sequence analysis of the chitin synthase A gene of the Dutch elm pathogen Ophiostoma novo-ulmi indicates a close association with the human pathogen Sporothrix schenckii. Gene 237:215–221. 98. Hirano, M., et al. 2006. A case of feline sporotrichosis. J. Vet. Med. Sci. 68:283–284. 99. Hiruma, M., T. Katoh, I. Yamamoto, and S. Kagawa. 1987. Local hyperthermia in the treatment of sporotrichosis. Mykosen 30:315–321. 100. Hogan, L. H., B. S. Klein, and S. M. Levitz. 1996. Virulence factors of medically important fungi. Clin. Microbiol. Rev. 9:469–488. 101. Howard, D. H., and G. F. Orr. 1963. Comparison of strains of Sporotrichum schenckii isolated from nature. J. Bacteriol. 85:816–821. 102. Howell, S. J., and J. S. Toohey. 1998. Sporotrichal arthritis in south central Kansas. Clin. Orthop. Relat. Res. 1998:207–214. 103. Ishizaki, H., et al. 2009. Mitochondrial DNA analysis of Sporothrix schenckii in India, Thailand, Brazil, Colombia, Guatemala and Mexico. Nippon Ishinkin Gakkai Zasshi 50:19–26.

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

651

104. Ishizaki, H., et al. 2004. Mitochondrial DNA analysis of Sporothrix schenckii from China, Korea and Spain. Nippon Ishinkin Gakkai Zasshi 45:23–25. 105. Itoh, M., S. Okamoto, and H. Kariya. 1986. Survey of 200 cases of sporotrichosis. Dermatologica 172:209–213. 106. Jacobson, E. S. 2000. Pathogenic roles for fungal melanins. Clin. Microbiol. Rev. 13:708–717. 107. Johnson, E. M., A. Szekely, and D. W. Warnock. 1998. In-vitro activity of voriconazole, itraconazole and amphotericin B against filamentous fungi. J. Antimicrob. Chemother. 42:741–745. 108. Kajiwara, H., M. Saito, S. Ohga, T. Uenotsuchi, and S. Yoshida. 2004. Impaired host defense against Sporothrix schenckii in mice with chronic granulomatous disease. Infect. Immun. 72:5073–5079. 109. Kanbe, T., et al. 2005. Rapid and specific identification of Sporothrix schenckii by PCR targeting the DNA topoisomerase II gene. J. Dermatol. Sci. 38:99–106. 110. Kano, R., Y. Nakamura, S. Watanabe, H. Tsujimoto, and A. Hasegawa. 2001. Identification of Sporothrix schenckii based on sequences of the chitin synthase 1 gene. Mycoses 44:261–265. 111. Kaplan, W., J. R. Broderson, and J. N. Pacific. 1982. Spontaneous systemic sporotrichosis in nine-banded armadillos (Dasypus novemcinctus). Sabouraudia 20:289–294. 112. Karlin, J. V., and H. S. Nielsen, Jr. 1970. Serologic aspects of sporotrichosis. J. Infect. Dis. 121:316–327. 113. Kashima, T., R. Honma, S. Kishi, and J. Hirato. 2010. Bulbar conjunctival sporotrichosis presenting as a salmon-pink tumor. Cornea 29:573–576. 114. Kauffman, C. A. 1999. Sporotrichosis. Clin. Infect. Dis. 29:231–237. 115. Kauffman, C. A., B. Bustamante, S. W. Chapman, and P. G. Pappas. 2007. Clinical practice guidelines for the management of sporotrichosis: 2007 update by the Infectious Diseases Society of America. Clin. Infect. Dis. 45:1255–1265. 116. Kazanas, N. 1986. Foodborne Sporothrix schenckii: infectivity for mice by intraperitoneal and intragastric inoculation with conidia. Mycopathologia 95:3–16. 117. Kazanas, N., and G. J. Jackson. 1983. Sporothrix schenckii isolated from edible black fungus mushrooms. J. Food Prot. 46:714–716. 118. Kenyon, E. M., L. H. Russell, and D. N. McMurray. 1984. Isolation of Sporothrix schenckii from potting soil. Mycopathologia 87:128. 119. Kikuchi, I., K. Morimoto, S. Kawana, and H. Tanuma. 2006. Usefulness of itraconazole for sporotrichosis in Japan: study of three cases and literature comparison of therapeutic effects before and after release on the market. Eur. J. Dermatol. 16:42–47. 120. Kohler, L. M., J. S. Hamdan, and T. C. Ferrari. 2007. Successful treatment of a disseminated Sporothrix schenckii infection and in vitro analysis for antifungal susceptibility testing. Diagn. Microbiol. Infect. Dis. 58:117–120. 121. Kohler, L. M., P. C. Monteiro, R. C. Hahn, and J. S. Hamdan. 2004. In vitro susceptibilities of isolates of Sporothrix schenckii to itraconazole and terbinafine. J. Clin. Microbiol. 42:4319–4320. 122. Kohler, L. M., B. M. Soares, D. de Assis Santos, M. E. Da Silva Barros, and J. S. Hamdan. 2006. In vitro susceptibility of isolates of Sporothrix schenckii to amphotericin B, itraconazole, and terbinafine: comparison of yeast and mycelial forms. Can. J. Microbiol. 52:843–847. 123. Kong, X., T. Xiao, J. Lin, Y. Wang, and H. D. Chen. 2006. Relationships among genotypes, virulence and clinical forms of Sporothrix schenckii infection. Clin. Microbiol. Infect. 12:1077–1081. 124. Kovarik, C. L., E. Neyra, and B. Bustamante. 2008. Evaluation of cats as the source of endemic sporotrichosis in Peru. Med. Mycol. 46:53–56. 125. Kumar, R., E. van der Smissen, and J. Jorizzo. 1984. Systemic sporotrichosis with osteomyelitis. J. Can. Assoc. Radiol. 35:83–84. 126. Kwon-Chung, K. J., and J. E. Bennet. 1992. Medical mycology. Lea & Febiger, Philadelphia, PA. 127. Kwon-Chung, K. J., W. B. Hill, and J. E. Bennett. 1981. New, special stain for histopathological diagnosis of cryptococcosis. J. Clin. Microbiol. 13:383– 387. 128. Lacaz, C. S., E. Porto, E. M. Heins-Vaccari, and N. T. Melo. 1998. Guia para identificac¸˜ao: fungos, actinomicetos e algas de interesse me´dico. Sarvier, Sa˜o Paulo, Brazil. 129. Lacaz, C. S., E. Porto, J. E. C. Martins, E. M. Heins-Vaccari, and N. T. Melo. 2002. Tratado de micologia me´dica. Sarvier, Sa˜o Paulo, Brazil. 130. Larone, D. H. 2002. Medically important fungi: a guide to identification, 4th ed. ASM Press, Washington, DC. 131. Latge, J. P. 1999. Aspergillus fumigatus and aspergillosis. Clin. Microbiol. Rev. 12:310–350. 132. Lavalle, P., and F. Mariat. 1983. Sporotrichosis. Bull. Inst. Pasteur 81:295– 322. 133. Lima, O. C., et al. 2004. Immunofluorescence and flow cytometry analysis of fibronectin and laminin binding to Sporothrix schenckii yeast cells and conidia. Microb. Pathog. 37:131–140. 134. Lima, O. C., et al. 1999. Adhesion of the human pathogen Sporothrix schenckii to several extracellular matrix proteins. Braz. J. Med. Biol. Res. 32:651–657. 135. Lima, O. C., et al. 2001. Involvement of fungal cell wall components in

652

136. 137.

138.

139. 140.

141. 142.

143.

144.

145. 146.

147.

148.

149. 150.

151. 152.

153.

154.

155.

156.

157.

158. 159.

160.

161. 162.

163.

164.

BARROS ET AL. adhesion of Sporothrix schenckii to human fibronectin. Infect. Immun. 69: 6874–6880. Lin, J., et al. 1999. Mitochondrial DNA analysis of Sporothrix schenckii clinical isolates from China. Mycopathologia 148:69–72. Lindsley, M. D., S. F. Hurst, N. J. Iqbal, and C. J. Morrison. 2001. Rapid identification of dimorphic and yeast-like fungal pathogens using specific DNA probes. J. Clin. Microbiol. 39:3505–3511. Lloyd, K. O., and M. A. Bitoon. 1971. Isolation and purification of a peptido-rhamnomannan from the yeast form of Sporothrix schenckii. Structural and immunochemical studies. J. Immunol. 107:663–671. Reference deleted. Loureiro y Penha, C. V., and L. M. Lopes Bezerra. 2000. Concanavalin A-binding cell wall antigens of Sporothrix schenckii: a serological study. Med. Mycol. 38:1–7. Lutz, A., and A. Splendore. 1907. Sobre uma mycose observada em homens e ratos. Rev. Med. Sa˜o Paulo 21:433–450. Lyon, G. M., et al. 2003. Population-based surveillance and a case-control study of risk factors for endemic lymphocutaneous sporotrichosis in Peru. Clin. Infect. Dis. 36:34–39. Mackinnon, J. E., I. A. Conti-Diaz, E. Gezuele, E. Civila, and S. da Luz. 1969. Isolation of Sporothrix schenckii from nature and considerations on its pathogenicity and ecology. Sabouraudia 7:38–45. Macotela-Ruiz, E., and E. Nochebuena-Ramos. 2006. Esporotricosis en algunas comunidades rurales de la sierra norte de Puebla: informe de 55 casos (Septiembre 1995-Diciembre 2005). Gac. Med. Mex. 142:377–380. Madrid, H., et al. 2009. Sporothrix globosa, a pathogenic fungus with widespread geographical distribution. Rev. Iberoam. Micol. 26:218–222. Magand, F., J. L. Perrot, F. Cambazard, M. H. Raberin, and B. Labeille. 2009. Sporotrichose cutane´e autochtone franc¸aise. Ann. Dermatol. Venereol. 136:273–275. Maia, D. C., M. F. Sassa, M. C. Placeres, and I. Z. Carlos. 2006. Influence of Th1/Th2 cytokines and nitric oxide in murine systemic infection induced by Sporothrix schenckii. Mycopathologia 161:11–19. Malliaris, S. D., J. N. Steenbergen, and A. Casadevall. 2004. Cryptococcus neoformans var. gattii can exploit Acanthamoeba castellanii for growth. Med. Mycol. 42:149–158. Manohar, A., and M. N. Nizlan. 2008. Chronic nonhealing ulcer of the right thumb with multiple subcutaneous nodules. Orthopedics 31:710. Marimon, R., et al. 2007. Sporothrix brasiliensis, S. globosa, and S. mexicana, three new Sporothrix species of clinical interest. J. Clin. Microbiol. 45:3198– 3206. Marimon, R., J. Gene, J. Cano, and J. Guarro. 2008. Sporothrix luriei: a rare fungus from clinical origin. Med. Mycol. 46:621–625. Marimon, R., C. Serena, J. Gene, J. Cano, and J. Guarro. 2008. In vitro antifungal susceptibilities of five species of Sporothrix. Antimicrob. Agents Chemother. 52:732–734. Masoko, P., L. K. Mdee, L. J. Mampuru, and J. N. Eloff. 2008. Biological activity of two related triterpenes isolated from Combretum nelsonii (Combretaceae) leaves. Nat. Prod. Res. 22:1074–1084. Masoko, P., J. Picard, R. L. Howard, L. J. Mampuru, and J. N. Eloff. 2010. In vivo antifungal effect of Combretum and Terminalia species extracts on cutaneous wound healing in immunosuppressed rats. Pharm. Biol. 48:621– 632. McGinnis, M. R., N. Nordoff, R. K. Li, L. Pasarell, and D. W. Warnock. 2001. Sporothrix schenckii sensitivity to voriconazole, itraconazole and amphotericin B. Med. Mycol. 39:369–371. Mehta, K. I., N. L. Sharma, A. K. Kanga, V. K. Mahajan, and N. Ranjan. 2007. Isolation of Sporothrix schenckii from the environmental sources of cutaneous sporotrichosis patients in Himachal Pradesh, India: results of a pilot study. Mycoses 50:496–501. Meinerz, A. R., et al. 2007. Suscetibilidade in vitro de isolados de Sporothrix schenckii frente `a terbinafina e itraconazol. Rev. Soc. Bras. Med. Trop. 40:60–62. Mendes-Giannini, M. J., et al. 2006. Binding of extracellular matrix proteins to Paracoccidioides brasiliensis. Microbes Infect. 8:1550–1559. Mendonca-Hagler, L. C., L. R. Travassos, K. O. Lloyd, and H. J. Phaff. 1974. Deoxyribonucleic acid base composition and hybridization studies on the human pathogen Sporothrix schenckii and Ceratocystis species. Infect. Immun. 9:934–938. Mendoza, M., P. Alvarado, E. Diaz de Torres, L. Lucena, and M. C. de Albornoz. 2005. Comportamiento fisiologico y de sensibilidad in vitro de aislamientos de Sporothrix schenckii mantenidos 18 an ˜os por do ´s metodos de preservacio ´n. Rev. Iberoam. Micol. 22:151–156. Mendoza, M., et al. 2002. Production of culture filtrates of Sporothrix schenckii in diverse culture media. Med. Mycol. 40:447–454. Mendoza, M., E. Diaz, P. Alvarado, E. Romero, and M. C. Bastardo de Albornoz. 2007. Aislamiento de Sporothrix schenckii del medio ambiente en Venezuela. Rev. Iberoam. Micol. 24:317–319. Mesa-Arango, A. C., et al. 2002. Phenotyping and genotyping of Sporothrix schenckii isolates according to geographic origin and clinical form of sporotrichosis. J. Clin. Microbiol. 40:3004–3011. Migaki, G., R. L. Font, W. Kaplan, and E. D. Asper. 1978. Sporotrichosis in

CLIN. MICROBIOL. REV.

165. 166. 167.

168.

169.

170. 171.

172.

173.

174.

175.

176. 177.

178.

179. 180. 181.

182.

183.

184. 185. 186. 187.

188. 189.

190. 191. 192.

193. 194.

195. 196.

a Pacific white-sided dolphin (Lagenorhynchus obliquidens). Am. J. Vet. Res. 39:1916–1919. Milby, A. H., N. D. Pappas, J. O’Donnell, and D. J. Bozentka. 2010. Sporotrichosis of the upper extremity. Orthopedics 16:273–275. Miller, S. D., and J. H. Keeling. 2002. Ant sting sporotrichosis. Cutis 69:439–442. Miranda, L. H., et al. 2010. Comparative histopathological study of sporotrichosis and American tegumentary leishmaniasis in dogs from Rio de Janeiro. J. Comp. Pathol. 143:1–7. Mora-Cabrera, M., R. A. Alonso, R. Ulloa-Arvizu, and H. Torres-Guerrero. 2001. Analysis of restriction profiles of mitochondrial DNA from Sporothrix schenckii. Med. Mycol. 39:439–444. Morera-Lopez, Y., J. M. Torres-Rodriguez, and T. Jimenez-Cabello. 2005. Estudio de la sensibilidad in vitro de aislamientos clínicos de mohos y levaduras a itraconazol y voriconazol. Rev. Iberoam. Micol. 22:105–109. Morris-Jones, R. 2002. Sporotrichosis. Clin. Exp. Dermatol. 27:427–431. Morris-Jones, R., et al. 2003. Synthesis of melanin-like pigments by Sporothrix schenckii in vitro and during mammalian infection. Infect. Immun. 71:4026–4033. Nakai, T., et al. 2003. In vitro antifungal activity of Micafungin (FK463) against dimorphic fungi: comparison of yeast-like and mycelial forms. Antimicrob. Agents Chemother. 47:1376–1381. Nascimento, R. C., and S. R. Almeida. 2005. Humoral immune response against soluble and fractionate antigens in experimental sporotrichosis. FEMS Immunol. Med. Microbiol. 43:241–247. Nascimento, R. C., et al. 2008. Passive immunization with monoclonal antibody against a 70-kDa putative adhesin of Sporothrix schenckii induces protection in murine sporotrichosis. Eur. J. Immunol. 38:3080–3089. Newton, P. N., W. H. Chung, R. Phetsouvanh, and N. J. White. 2005. Sporotrichosis, Plain of Jars, Lao People’s Democratic Republic. Emerg. Infect. Dis. 11:1496–1497. Nobre, M. O., et al. 2005. Differences in virulence between isolates of feline sporotrichosis. Mycopathologia 160:43–49. Noriega, C. T., R. R. Garay, G. Sabanero, R. T. Basurto, and M. SabaneroLopez. 1993. Sporothrix schenckii: culturas en diferentes suelos. Rev. Latinoam. Micol. 35:191–194. Odabasi, Z., V. L. Paetznick, J. R. Rodriguez, E. Chen, and L. OstroskyZeichner. 2004. In vitro activity of anidulafungin against selected clinically important mold isolates. Antimicrob. Agents Chemother. 48:1912–1915. Oliveira, M. M. E. 2009. M.Sc. thesis. Fundac¸˜ao Oswaldo Cruz, Rio de Janeiro, Brazil. Oliveira, M. M. E., et al. 2010. Sporotrichosis caused by Sporothrix globosa in Rio de Janeiro, Brazil: case report. Mycopathologia 169:359–363. O’Reilly, L. C., and S. A. Altman. 2006. Macrorestriction analysis of clinical and environmental isolates of Sporothrix schenckii. J. Clin. Microbiol. 44: 2547–2552. Orellana, A., G. Moreno-Coutino, E. Poletti, M. E. Vega, and R. Arenas. 2009. Esporotricosis fija con cuerpo asteroide junto al fragmento vegetal. Rev. Iberoam. Micol. 26:250–251. Palomino, J., O. Saeed, P. Daroca, and J. Lasky. 2009. A 47-year-old man with cough, dyspnea, and an abnormal chest radiograph. Chest 135:872– 875. Pappas, P. G., et al. 2000. Sporotrichosis in Peru: description of an area of hyperendemicity. Clin. Infect. Dis. 30:65–70. Powell, K. E., et al. 1978. Cutaneous sporotrichosis in forestry workers. Epidemic due to contaminated Sphagnum moss. JAMA 240:232–235. Pupo, J. A. 1917. Frequencia da sporotrichose em Sa˜o Paulo. Ann. Paulistas Med. Cirurgia 8:53–68. Quintella, L. P., S. R. Lambert Passos, A. C. Francesconi do Vale, et al. 2011. Histopathology of cutaneous sporotrichosis in Rio de Janeiro: a series of 119 consecutive cases. J. Cutan. Pathol. 38:25–32. Ramesh, V. 2007. Sporotrichoid cutaneous tuberculosis. Clin. Exp. Dermatol. 32:680–682. Ramirez, J., R. P. Byrd, Jr., and T. M. Roy. 1998. Chronic cavitary pulmonary sporotrichosis: efficacy of oral itraconazole. J. Ky. Med. Assoc. 96:103– 105. Ramos-e-Silva, M., C. Vasconcelos, S. Carneiro, and T. Cestari. 2007. Sporotrichosis. Clin. Dermatol. 25:181–187. Read, S. I., and L. C. Sperling. 1982. Feline sporotrichosis. Transmission to man. Arch. Dermatol. 118:429–431. Reis, R. S., et al. 2009. Molecular characterisation of Sporothrix schenckii isolates from humans and cats involved in the sporotrichosis epidemic in Rio de Janeiro, Brazil. Mem. Inst. Oswaldo Cruz 104:769–774. Reiss, E. 1986. Molecular immunology of mycotic and actinomycotic infections. Elsevier, Amsterdam, The Netherlands. Reiss, E., T. Obayashi, K. Orle, M. Yoshida, and R. M. Zancope-Oliveira. 2000. Non-culture based diagnostic tests for mycotic infections. Med. Mycol. 38(Suppl. 1):147–159. Resto, S., and N. Rodriguez-del Valle. 1988. Yeast cell cycle of Sporothrix schenckii. J. Med. Vet. Mycol. 26:13–24. Rippon, J. 1988. Sporotrichosis, p. 325–352. In J. Rippon (ed.), Medical

VOL. 24, 2011

197.

198.

199.

200.

201.

202.

203.

204.

205.

206.

207.

208.

209. 210.

211.

212.

213.

214. 215.

216.

217. 218. 219.

220. 221.

222. 223.

mycology—the pathogenic fungi and the pathogenic actinomycetes, 3rd ed. W. B. Saunders Company, Philadelphia, PA. Rodrigues, M. L., L. Nimrichter, D. L. Oliveira, J. D. Nosanchuk, and A. Casadevall. 2008. Vesicular trans-cell wall transport in fungi: a mechanism for the delivery of virulence-associated macromolecules? Lipid Insights 2:27–40. Rodrigues, M. T., and M. A. Resende. 1996. Epidemiologic skin test survey of sensitivity to paracoccidioidin, histoplasmin and sporotrichin among gold mine workers of Morro Velho Mining, Brazil. Mycopathologia 135:89–98. Rodriguez-del Valle, N., and J. R. Rodriguez-Medina. 1993. Calcium stimulates molecular and cellular events during the yeast-to-mycelium transition in Sporothrix schenckii. J. Med. Vet. Mycol. 31:43–53. Rodriguez-Del Valle, N., M. Rosario, and G. Torres-Blasini. 1983. Effects of pH, temperature, aeration and carbon source on the development of the mycelial or yeast forms of Sporothrix schenckii from conidia. Mycopathologia 82:83–88. Rodriguez-Tudela, J. L., et al. 2008. Method for the determination of broth dilution minimum inhibitory concentrations of antifungal agents for conidia forming moulds. EUCAST E.DEF. 9.1. Romero-Martinez, R., M. Wheeler, A. Guerrero-Plata, G. Rico, and H. Torres-Guerrero. 2000. Biosynthesis and functions of melanin in Sporothrix schenckii. Infect. Immun. 68:3696–3703. Rosa, W. D., E. Gezuele, L. Calegari, and F. Goni. 2008. Asteroid body in sporotrichosis. Yeast viability and biological significance within the host immune response. Med. Mycol. 46:443–448. Rosa, W. D., E. Gezuele, L. Calegari, and F. Gon ˜ i. 2009. Excretion-secretion products and proteases from live Sporothrix schenckii yeast phase: immunological detection and cleavage of human IgG. Rev. Inst. Med. Trop. Sa˜o Paulo 51:1–7. Rubio, G., G. Sanchez, L. Porras, and Z. Alvarado. 2010. Esporotricosis: prevalencia, perfil clinico y epidemiologico en un centro de referencia en Colombia. Rev. Iberoam. Micol. 27:75–79. Ruiz-Baca, E., et al. 2009. Isolation and some properties of a glycoprotein of 70 kDa (Gp70) from the cell wall of Sporothrix schenckii involved in fungal adherence to dermal extracellular matrix. Med. Mycol. 47:185–196. Ruiz-Cendoya, M., M. M. Rodriguez, M. Marine, F. J. Pastor, and J. Guarro. 2008. In vitro interactions of itraconazole and micafungin against clinically important filamentous fungi. Int. J. Antimicrob. Agents 32:418– 420. Safdar, A. 2003. Clinical microbiological case: infection imitating lymphocutaneous sporotrichosis during pregnancy in a healthy woman from the south-eastern USA. Clin. Microbiol. Infect. 9:221, 224–226. Salaki, J. S., D. B. Louria, and H. Chmel. 1984. Fungal and yeast infections of the central nervous system. A clinical review. Medicine 63:108–132. Sanchez-Aleman, M. A., J. Araiza, and A. Bonifaz. 2004. Aislamiento y caracterizacion de cepas silvestres de Sporothrix schenckii e investigacion de reactores a la esporotricina. Gac. Med. Mex. 140:507–512. Sandhu, G. S., B. C. Kline, L. Stockman, and G. D. Roberts. 1995. Molecular probes for diagnosis of fungal infections. J. Clin. Microbiol. 33:2913– 2919. Saravanakumar, P. S., P. Eslami, and F. A. Zar. 1996. Lymphocutaneous sporotrichosis associated with a squirrel bite: case report and review. Clin. Infect. Dis. 23:647–648. Sashidhara, K. V., S. P. Singh, and P. K. Shukla. 2009. Antimicrobial evaluation of clerodane diterpenes from Polyalthia longifolia var. pendula. Nat. Prod. Commun. 4:327–330. Scheinfeld, N. 2007. A review of the new antifungals: posaconazole, micafungin, and anidulafungin. J. Drugs Dermatol. 6:1249–1251. Schell, W. 1998. Agents of chromoblastomycosis and sporotrichosis, p. 315–336. In L. Ajello and R. J. Hay (ed.), Topley & Wilson’s microbiology and microbial infections, 9th ed., vol. 4. Arnold, London, United Kingdom. Schell, W. A., I. F. Salkin, L. Pasarell, and M. R. McGinnis. 1999. Bipolaris, Exophiala, Scedosporium, Sporothrix and other dematiaceous fungi, p. 1295– 1317. In P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (ed.), Manual of clinical microbiology, 7th ed. ASM Press, Washington, DC. Schenck, B. R. 1898. On refractory subcutaneous abscess caused by a fungus possibly related to the Sporotricha. Bull. Johns Hopkins Hosp. 9:286–290. Schubach, A., M. B. Barros, and B. Wanke. 2008. Epidemic sporotrichosis. Curr. Opin. Infect. Dis. 21:129–133. Schubach, A., M. B. de Lima Barros, T. M. Schubach, A. C. Francesconido-Valle, et al. 2005. Primary conjunctival sporotrichosis: two cases from a zoonotic epidemic in Rio de Janeiro, Brazil. Cornea 24:491–493. Schubach, A. O., T. M. Schubach, and M. B. Barros. 2005. Epidemic cat-transmitted sporotrichosis. N. Engl. J. Med. 353:1185–1186. Schubach, T. M., A. de Oliveira Schubach, R. S. dos Reis, et al. 2002. Sporothrix schenckii isolated from domestic cats with and without sporotrichosis in Rio de Janeiro, Brazil. Mycopathologia 153:83–86. Schubach, T. M., et al. 2004. Evaluation of an epidemic of sporotrichosis in cats: 347 cases (1998-2001). J. Am. Vet. Med. Assoc. 224:1623–1629. Schubach, T. M., et al. 2006. Canine sporotrichosis in Rio de Janeiro,

SPOROTHRIX SCHENCKII AND SPOROTRICHOSIS

224.

225. 226. 227.

228.

229. 230.

231.

232.

233.

234. 235.

236.

237.

238.

239. 240.

241. 242. 243.

244. 245.

246.

247.

248.

249.

250.

251.

252.

653

Brazil: clinical presentation, laboratory diagnosis and therapeutic response in 44 cases (1998-2003). Med. Mycol. 44:87–92. Schubach, T. M., et al. 2003. Haematogenous spread of Sporothrix schenckii in cats with naturally acquired sporotrichosis. J. Small Anim. Pract. 44:395– 398. Schubach, T. M., et al. 2003. Pathology of sporotrichosis in 10 cats in Rio de Janeiro. Vet. Rec. 152:172–175. Schubach, T. M., et al. 2001. Isolation of Sporothrix schenckii from the nails of domestic cats (Felis catus). Med. Mycol. 39:147–149. Schwartz, D. A. 1989. Sporothrix tenosynovitis—differential diagnosis of granulomatous inflammatory disease of the joints. J. Rheumatol. 16:550– 553. Scott, E. N., L. Kaufman, A. C. Brown, and H. G. Muchmore. 1987. Serologic studies in the diagnosis and management of meningitis due to Sporothrix schenckii. N. Engl. J. Med. 317:935–940. Scott, E. N., and H. G. Muchmore. 1989. Immunoblot analysis of antibody responses to Sporothrix schenckii. J. Clin. Microbiol. 27:300–304. Scott, E. N., H. G. Muchmore, and D. P. Fine. 1986. Activation of the alternative complement pathway by Sporothrix schenckii. Infect. Immun. 51:6–9. Sgarbi, D. B., et al. 1997. Isolation of ergosterol peroxide and its reversion to ergosterol in the pathogenic fungus Sporothrix schenckii. Mycopathologia 139:9–14. Sia, R. A., K. B. Lengeler, and J. Heitman. 2000. Diploid strains of the pathogenic basidiomycete Cryptococcus neoformans are thermally dimorphic. Fungal Genet. Biol. 29:153–163. Sigler, L., et al. 1990. Microbiology and potential virulence of Sporothrix cyanescens, a fungus rarely isolated from blood and skin. J. Clin. Microbiol. 28:1009–1015. Silva-Vergara, M. L., et al. 2005. Multifocal sporotrichosis with meningeal involvement in a patient with AIDS. Med. Mycol. 43:187–190. Silveira, C. P., et al. 2009. MICs and minimum fungicidal concentrations of amphotericin B, itraconazole, posaconazole and terbinafine in Sporothrix schenckii. J. Med. Microbiol. 58:1607–1610. Singer, J. I., and J. E. Muncie. 1952. Sporotrichosis. Etiologic considerations and report of additional cases from New York. N. Y. State J. Med. 52:2147–2153. Smith, M. T., and W. H. Batenburg-Van der Vegte. 1985. Ultrastructure in septa in Blastobotrys and Sporothrix. Antonie Van Leeuwenhoek 51:121– 128. Song, Y., et al. 2011. Efficacy and safety of itraconazole pulses vs. continuous regimen in cutaneous sporotrichosis. J. Eur. Acad. Dermatol. Venereol. 25:302–305. Stalkup, J. R., K. Bell, and T. Rosen. 2002. Disseminated cutaneous sporotrichosis treated with itraconazole. Cutis 69:371–374. Steenbergen, J. N., J. D. Nosanchuk, S. D. Malliaris, and A. Casadevall. 2004. Interaction of Blastomyces dermatitidis, Sporothrix schenckii, and Histoplasma capsulatum with Acanthamoeba castellanii. Infect. Immun. 72: 3478–3488. Stevens, D. A. 2002. Diagnosis of fungal infections: current status. J. Antimicrob. Chemother. 49(Suppl. 1):11–19. St-Germain, G., and R. C. Summerbell. 1996. Identifying filamentous fungi. A clinical laboratory handbook. Star Publishing Company, Belmont, CA. Suzuki, K., M. Kawasaki, and H. Ishizaki. 1988. Analysis of restriction profiles of mitochondrial DNA from Sporothrix schenckii and related fungi. Mycopathologia 103:147–151. Svoboda, A., and A. Trujillo-Gonzalez. 1990. Sporothrix schenckii—a freezefracture study. J. Basic Microbiol. 30:371–378. Taboada, J. 2000. Systemic mycoses, p. 453–476. In S. Ettinger and E. Feldman (ed.), Textbook of veterinary internal medicine—diseases of the dog and cat, 5th ed., vol. 1. W. B. Saunders Company, Philadelphia, PA. Tachibana, T., T. Matsuyama, and M. Mitsuyama. 1998. Characteristic infectivity of Sporothrix schenckii to mice depending on routes of infection and inherent fungal pathogenicity. Med. Mycol. 36:21–27. Tachibana, T., T. Matsuyama, and M. Mitsuyama. 1999. Involvement of CD4⫹ T cells and macrophages in acquired protection against infection with Sporothrix schenckii in mice. Med. Mycol. 37:397–404. Takenaka, M., S. Sato, and K. Nishimoto. 2009. Survey of 155 sporotrichosis cases examined in Nagasaki Prefecture from 1951 to 2007. Nippon Ishinkin Gakkai Zasshi 50:101–108. Tateishi, T., S. Y. Murayama, F. Otsuka, and H. Yamaguchi. 1996. Karyotyping by PFGE of clinical isolates of Sporothrix schenckii. FEMS Immunol. Med. Microbiol. 13:147–154. Teixeira, P. A., et al. 2010. L-DOPA accessibility in culture medium increases melanin expression and virulence of Sporothrix schenckii yeast cells. Med. Mycol. 48:687–695. Teixeira, P. A., et al. 2009. Cell surface expression of adhesins for fibronectin correlates with virulence in Sporothrix schenckii. Microbiology 155:3730– 3738. Tlougan, B. E., J. O. Podjasek, S. P. Patel, X. H. Nguyen, and R. C. Hansen. 2009. Neonatal sporotrichosis. Pediatr. Dermatol. 26:563–565.

654

BARROS ET AL.

CLIN. MICROBIOL. REV.

253. Tobin, E. H., and W. W. Jih. 2001. Sporotrichoid lymphocutaneous infections: etiology, diagnosis and therapy. Am. Fam. Physician 63:326–332. 254. Toledo, M. S., et al. 2010. Effect of anti-glycosphingolipid monoclonal antibodies in pathogenic fungal growth and differentiation. Characterization of monoclonal antibody MEST-3 directed to Manpa133Manpa32IPC. BMC Microbiol. 10:47. 255. Torinuki, W., and H. Tagami. 1985. Complement activation by Sporothrix schenckii. Arch. Dermatol. Res. 277:332–333. 256. Torres-Guerrero, H. 1999. Ploidy studies in Sporothrix schenckii. Fungal Genet. Biol. 27:49–54. 257. Trilles, L., et al. 2005. In vitro antifungal susceptibilities of Sporothrix schenckii in two growth phases. Antimicrob. Agents Chemother. 49:3952– 3954. 258. Uenotsuchi, T., et al. 2006. Differential induction of Th1-prone immunity by human dendritic cells activated with Sporothrix schenckii of cutaneous and visceral origins to determine their different virulence. Int. Immunol. 18: 1637–1646. 259. Valentin-Berrios, S., W. Gonzalez-Velazquez, L. Perez-Sanchez, R. Gonzalez-Mendez, and N. Rodriguez-Del Valle. 2009. Cytosolic phospholipase A2: a member of the signalling pathway of a new G protein alpha subunit in Sporothrix schenckii. BMC Microbiol. 9:100. 260. Valle-Aviles, L., S. Valentin-Berrios, R. R. Gonzalez-Mendez, and N. Rodriguez-Del Valle. 2007. Functional, genetic and bioinformatic characterization of a calcium/calmodulin kinase gene in Sporothrix schenckii. BMC Microbiol. 7:107. 261. van Duin, D., A. Casadevall, and J. D. Nosanchuk. 2002. Melanization of Cryptococcus neoformans and Histoplasma capsulatum reduces their susceptibilities to amphotericin B and caspofungin. Antimicrob. Agents Chemother. 46:3394–3400.

262. Vicentini, A. P., et al. 1994. Binding of Paracoccidioides brasiliensis to laminin through surface glycoprotein gp43 leads to enhancement of fungal pathogenesis. Infect. Immun. 62:1465–1469. 263. Vilela, R., G. F. Souza, G. Fernandes Cota, and L. Mendoza. 2007. Cutaneous and meningeal sporotrichosis in a HIV patient. Rev. Iberoam. Micol. 24:161–163. 264. Wenker, C. J., L. Kaufman, L. N. Bacciarini, and N. Robert. 1998. Sporotrichosis in a nine-banded armadillo (Dasypus novemcinctus). J. Zoo Wildl. Med. 29:474–478. 265. Woods, J. P. 2003. Knocking on the right door and making a comfortable home: Histoplasma capsulatum intracellular pathogenesis. Curr. Opin. Microbiol. 6:327–331. 266. Xu, T. H., et al. 2010. Identification of Sporothrix schenckii of various mtDNA types by nested PCR assay. Med. Mycol. 48:161–165. 267. Xue, S. L., and L. Li. 2009. Oral potassium iodide for the treatment of sporotrichosis. Mycopathologia 167:355–356. 268. Yang, D. J., et al. 2006. Disseminated sporotrichosis mimicking sarcoidosis. Int. J. Dermatol. 45:450–453. 269. Yegneswaran, P. P., et al. 2009. Zoonotic sporotrichosis of lymphocutaneous type in a man acquired from a domesticated feline source: report of a first case in southern Karnataka, India. Int. J. Dermatol. 48:1198–1200. 270. Zarnowski, R., and J. P. Woods. 2005. Glutathione-dependent extracellular ferric reductase activities in dimorphic zoopathogenic fungi. Microbiology 151:2233–2240. 271. Zhang, X., and J. H. Andrews. 1993. Evidence for growth of Sporothrix schenckii on dead but not on living sphagnum moss. Mycopathologia 123: 87–94.

Mo ˆnica Bastos de Lima Barros obtained her medical degree from the Federal University of the State of Rio de Janeiro, Brazil, in 1984. She completed her residency in public health at Oswaldo Cruz Foundation and subsequently her M.Sc. and Ph.D. degrees in infectious diseases from the Federal University of Rio de Janeiro. Her thesis was on sporotrichosis and the first description of an epidemic due to zoonotic transmission. During the past 10 years she has gained broad experience in the clinical aspects, epidemiology, treatment, and prevention of sporotrichosis. She works as a researcher and professor at the Oswaldo Cruz Foundation, affiliated with the Ministry of Health, developing studies in infectious diseases.

Armando de Oliveira Schubach obtained his medical degree from the University of the State of Rio de Janeiro, Brazil, in 1980, a specialization in dermatology from the Federal University of the State of Rio de Janeiro in 1990, an M.Sc. degree in tropical medicine from the Oswaldo Cruz Foundation (Fiocruz) in 1990, and a Ph.D. degree in parasite biology from Fiocruz in 1997. He is currently a senior researcher at the Evandro Chagas Clinical Research Institute (IPEC/Fiocruz), coordinator of the Laboratory of Leishmaniasis Surveillance at IPEC/Fiocruz, coordinator of the Postgraduation Program in Clinical Research in Infectious Diseases of IPEC/Fiocruz, Level 2 researcher of the National Council of Scientific and Technological Development (CNPq), leader of the Parasitic Diseases (CNPq) research group, and a “Scientist of Our State” from the Foundation for Research of Rio de Janeiro (FAPERJ). He has experience in the study of infectious and parasitic diseases, mainly the clinical, diagnostic, treatment, and epidemiological aspects of sporotrichosis and leishmaniasis.

Rodrigo Almeida Paes obtained his undergraduate degree from the Federal University of the State of Rio de Janeiro, Brazil, in 2004. Since 2002 he has worked at the Oswaldo Cruz Foundation, where he completed his M.Sc. degree in cellular and molecular biology in 2007. He is now a Ph.D. student, working on several aspects of S. schenckii virulence. He is also the supervisor of the Mycological Diagnosis section of the Mycology Laboratory of the Oswaldo Cruz Foundation and teaches medical mycology to technicians and undergraduate students.

CLINICAL MICROBIOLOGY REVIEWS, Oct. 2011, p. 655–681 0893-8512/11/$12.00 doi:10.1128/CMR.00005-11 Copyright © 2011, American Society for Microbiology. All Rights Reserved.

Vol. 24, No. 4

Trypanosoma cruzi and Chagas’ Disease in the United States Caryn Bern,1* Sonia Kjos,2 Michael J. Yabsley,3 and Susan P. Montgomery1 Division of Parasitic Diseases and Malaria, Center for Global Health, Centers for Disease Control and Prevention, Atlanta, Georgia1; Marshfield Clinic Research Foundation, Marshfield, Wisconsin2; and Department of Population Health, College of Veterinary Medicine, University of Georgia, Athens, Georgia3 INTRODUCTION .......................................................................................................................................................656 TRYPANOSOMA CRUZI LIFE CYCLE AND TRANSMISSION ..........................................................................656 Life Cycle .................................................................................................................................................................656 Transmission Routes ..............................................................................................................................................657 Vector-borne transmission.................................................................................................................................657 Congenital transmission ....................................................................................................................................657 Blood-borne transmission..................................................................................................................................657 Organ-derived transmission ..............................................................................................................................657 Oral transmission ...............................................................................................................................................657 TRIATOMINE VECTOR BIOLOGY AND ECOLOGY ........................................................................................657 Background ..............................................................................................................................................................657 Triatomine Distribution in the United States ....................................................................................................658 Description of U.S. Triatomine Species...............................................................................................................659 Triatoma gerstaeckeri (Stål) ................................................................................................................................659 Triatoma incrassata Usinger...............................................................................................................................660 Triatoma indictiva Neiva .....................................................................................................................................660 Triatoma lecticularia (Stål) .................................................................................................................................660 Triatoma neotomae Neiva....................................................................................................................................661 Triatoma protracta (Uhler) .................................................................................................................................661 Triatoma recurva (Stål).......................................................................................................................................661 Triatoma rubida (Uhler) .....................................................................................................................................661 Triatoma rubrofasciata (DeGeer) .......................................................................................................................661 Triatoma sanguisuga (Leconte) ..........................................................................................................................662 Paratriatoma hirsuta Barber...............................................................................................................................662 Human-Vector Interactions and T. cruzi Transmission Potential in the United States...............................662 ANIMAL RESERVOIRS OF TRYPANOSOMA CRUZI..........................................................................................663 Background ..............................................................................................................................................................663 Wildlife Reservoirs of T. cruzi in the United States ..........................................................................................663 Domestic and Exotic Animal Infections in the United States..........................................................................665 Canine Chagas’ disease......................................................................................................................................665 Primates and other exotic animals...................................................................................................................666 MOLECULAR EPIDEMIOLOGY OF T. CRUZI ...................................................................................................666 General Molecular Epidemiology .........................................................................................................................666 T. cruzi Genotypes in the United States ..............................................................................................................667 CLINICAL ASPECTS OF CHAGAS’ DISEASE.....................................................................................................668 Acute T. cruzi Infection ..........................................................................................................................................668 Congenital T. cruzi Infection .................................................................................................................................668 Chronic T. cruzi Infection ......................................................................................................................................668 Indeterminate form of chronic T. cruzi infection ...........................................................................................668 Cardiac Chagas’ disease ....................................................................................................................................668 Digestive Chagas’ disease ..................................................................................................................................668 T. cruzi Infection in the Immunocompromised Host .........................................................................................669 Acute T. cruzi infection in organ transplantation recipients........................................................................669 Reactivation of chronic T. cruzi infection in organ recipients......................................................................669 Reactivation Chagas’ disease in HIV/AIDS patients .....................................................................................669 DIAGNOSIS ................................................................................................................................................................669 Diagnosis of Acute T. cruzi Infection ...................................................................................................................669 Diagnosis of Congenital T. cruzi Infection ..........................................................................................................669 Diagnosis of Chronic T. cruzi infection ...............................................................................................................670

* Corresponding author. Mailing address: Division of Parasitic Diseases and Malaria, Center for Global Health, Centers for Disease Control and Prevention, 1600 Clifton Rd. N.E., Atlanta, GA 30333. Phone: (404) 718-4726. Fax: (404) 718-4816. E-mail: [email protected]. 655

656

BERN ET AL.

CLIN. MICROBIOL. REV.

Utility of PCR for Diagnosis or Monitoring .......................................................................................................670 TREATMENT..............................................................................................................................................................670 Antitrypanosomal Drugs ........................................................................................................................................670 Treatment of Acute and Congenital T. cruzi infection.......................................................................................670 Treatment of Chronic T. cruzi Infection ..............................................................................................................670 Management of the Immunocompromised Host ................................................................................................671 EPIDEMIOLOGY OF CHAGAS’ DISEASE ...........................................................................................................671 HUMAN CHAGAS’ DISEASE IN THE UNITED STATES..................................................................................671 Autochthonous Transmission to Humans ...........................................................................................................671 Chagas’ Disease Burden among Latin American Immigrants .........................................................................672 Blood-Borne Transmission and Blood Donor Screening ..................................................................................672 Organ Donor-Derived Transmission and Organ Donor Screening.................................................................673 Unanswered Questions and Priorities for Research and Programs................................................................673 REFERENCES ............................................................................................................................................................674 TRYPANOSOMA CRUZI LIFE CYCLE AND TRANSMISSION

INTRODUCTION Chagas’ disease is caused by the protozoan parasite Trypanosoma cruzi (234). World Health Organization disease burden estimates place Chagas’ disease first among parasitic diseases in the Americas, accounting for nearly 5 times as many disability-adjusted life years lost as malaria (343). An estimated 8 million people are currently infected, and 20 to 30% of these will develop symptomatic, potentially life-threatening Chagas’ disease (Table 1) (214). T. cruzi is carried in the guts of hematophagous triatomine bugs; transmission occurs when infected bug feces contaminate the bite site or intact mucous membranes. T. cruzi can also be transmitted through transfusion, through transplant, and congenitally (177, 234). Historically, transmission and morbidity were concentrated in rural areas of Latin America where poor housing conditions favor vector infestation. However, in the last several decades, successful vector control programs have substantially decreased transmission in rural areas, and migration has brought infected individuals to cities both within and outside Latin America (87, 111, 196). Since 1991, several subregional initiatives have made major advances in decreasing vector infestation in human dwellings and extending screening of the blood supply for T. cruzi (87, 269). In 2007, control efforts in Latin America were formally joined by an initiative to address the “globalization” of Chagas’ disease, recognizing the increasing presence of imported cases in Europe, North America, and Japan and the potential for local transmission through nonvectorial routes (344). The United States occupies an ambiguous position in this new initiative. While the United States has never participated in Latin American Chagas’ disease control programs, it cannot be classified as an area where the disease is “not endemic” in the same sense as Europe or Japan. The southern tier of states from Georgia to California contains established enzootic cycles of T. cruzi, involving several triatomine vector species and mammalian hosts such as raccoons, opossums, and domestic dogs (26, 151, 345). Nevertheless, most T. cruzi-infected individuals in the United States are immigrants from areas of endemicity in Latin America (29). This article will present an overview of clinical and epidemiological aspects of Chagas’ disease, with a focus on data and issues specific to T. cruzi and Chagas’ disease in the United States. Topics to be covered include vector biology and ecology, animal reservoirs, T. cruzi strain typing, human Chagas’ disease, and future research needed for control of Chagas’ disease in the United States.

Life Cycle Nearly all the salient features of the T. cruzi life cycle were described by Carlos Chagas, the scientist who discovered the organism, in 1909 (62). T. cruzi is a kinetoplastid protozoan which infects vertebrate and invertebrate hosts during defined stages in its life cycle (234, 292). The triatomine vector ingests circulating trypomastigotes when it takes a blood meal from an infected mammalian host. In the midgut of the vector, trypomastigotes transform through an intermediate form sometimes

TABLE 1. Countries where Chagas’ disease is endemic and estimates of the seroprevalence and number of infected inhabitants Region

Country where Chagas’ disease is endemica

Estimated seroprevalence (%)b

Estimated no. of infected individualsb

North America

United States Mexico

NDA 1.03

300,167c 1,100,000

Central America

Belize Costa Rica El Salvador Honduras Guatemala Nicaragua Panama

0.74 0.53 3.37 3.05 1.98 1.14 0.01

2,000 23,000 232,000 220,000 250,000 58,600 21,000

South America

Argentina Bolivia Brazil Chile Colombia Ecuador Guyana Suriname French Guiana Paraguay Peru Uruguay Venezuela

4.13 6.75 1.02 0.99 0.96 1.74 1.29 NDA NDA 2.54 0.69 0.66 1.16

1,600,000 620,000 1,900,000 160,200 436,000 230,000 18,000 NDA NDA 150,000 192,000 21,700 310,000

a Vector-borne T. cruzi transmission occurs, or occurred until recently, in parts of these countries. b Disease burden estimates are for the year 2005, based on references 29 and 214. NDA, No data available. c The number for the United States reflects the estimated number of infected immigrants from countries in Latin America where the disease is endemic. No estimate of the number of locally acquired infections is currently available.

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

657

called a spheromastigote to epimastigotes, the main replicating stage in the invertebrate host. Epimastigotes migrate to the hindgut and differentiate into infective metacyclic trypomastigotes, which are excreted with the feces of the vector. Metacyclic trypomastigotes enter through the bite wound or intact mucous membrane of the mammalian host and invade many types of nucleated cells through a lysosome-mediated mechanism (50). In the cytoplasm, trypomastigotes differentiate into the intracellular amastigote form, which replicates with a doubling time of about 12 h over a period of 4 to 5 days. At the end of this period, the amastigotes transform into trypomastigotes, the host cell ruptures, and the trypomastigotes are released into the circulation. The circulating parasites can then invade new cells and initiate new replicative cycles, and they are available to infect vectors that feed on the host. In the absence of successful antitrypanosomal treatment, the infection lasts for the lifetime of the mammalian host.

been documented in the literature (13 kidney, 1 kidney and pancreas, 3 liver, and 2 heart transplants) (16, 61, 66, 79, 99, 101, 157, 238, 279). The risk from heart transplantation is thought to be higher than that from kidney or liver transplantation (65). One case of transmission through unrelated cord blood transplantation has been reported (104). Oral transmission. Recently, increasing attention has focused on the oral route of T. cruzi transmission; several outbreaks attributed to contaminated fruit or sugar cane juice have been reported from Brazil and Venezuela (28, 82, 208). Most outbreaks are small, often affecting family groups in the Amazon region, where the palm fruit ac¸aí is a dietary staple that appears to be particularly vulnerable to contamination, perhaps from infected vectors living in the trees themselves (74, 208). The largest reported outbreak to date led to more than 100 infections among students and staff at a school in Caracas; locally prepared guava juice was implicated (82).

Transmission Routes

TRIATOMINE VECTOR BIOLOGY AND ECOLOGY

Vector-borne transmission. The vector-borne transmission route, occurring exclusively in the Americas, is still the predominant mechanism for new human infections. The feces of infected bugs contain metacyclic trypomastigotes that can enter the human body through the bite wound or through intact conjunctiva or other mucous membranes. Congenital transmission. Between 1 and 10% of infants of T. cruzi-infected mothers are born with congenital Chagas’ disease (14, 24, 289). Congenital transmission can occur from women themselves infected congenitally, perpetuating the disease in the absence of the vector (263). Factors reported to increase risk include higher maternal parasitemia level, less robust anti-T. cruzi immune responses, younger maternal age, HIV and, in an animal model, parasite strain (9, 32, 34, 107, 289). Blood-borne transmission. Transfusional T. cruzi transmission was postulated in 1936 and first documented in 1952 (109, 307). The risk of T. cruzi transmission per infected unit transfused is estimated to be 10 to 25%; platelet transfusions are thought to pose a higher risk than other components such as packed red cells (31, 308). In 1991, the prevalence of T. cruzi infection in donated blood units ranged from 1 to 60% in Latin American cities (268). Since then, blood donation screening has become accepted as an important pillar of the Chagas’ disease control initiatives (220, 269). Serological screening of blood components for T. cruzi is now compulsory in all but one of the countries in Latin America where the disease is endemic, and the prevalence of infection in screened donors has decreased substantially (196, 269). Nevertheless, Chagas’ disease screening coverage by country was estimated to vary from 25% to 100% in 2002, and the risk of transmission, though much decreased, has not been eliminated (269). The residual risk in Latin America where screening has been implemented is estimated to be 1:200,000 units (269, 308). Organ-derived transmission. Uninfected recipients who receive an organ from a T. cruzi-infected donor may develop acute T. cruzi infection. However, transmission is not universal; in a series of 16 uninfected recipients of kidneys from infected donors, only 3 (19%) acquired T. cruzi infection (238). Nineteen instances of transmission by organ transplantation have

Background The epidemiology of vector-borne T. cruzi is closely linked to the biological and ecological characteristics of local vectors and mammalian reservoir hosts. Triatomines of both sexes must take blood meals to develop through their nymphal stages to adults, and females require a blood meal to lay eggs. Thus, nymphs and adults of either sex may be infected with T. cruzi, but infection rates increase with increasing vector stage and age. Most domestic triatomine species feed nocturnally and are able to complete their blood meal without waking the host (169). The major Latin American vectors defecate during or immediately after taking a blood meal. T. cruzi infection is transmitted to wild mammals by sylvatic triatomine species; these bugs often colonize the nests of rodent or marsupial reservoir hosts (169, 311). Sylvatic triatomine adults may fly into human dwellings because of attraction by light and cause sporadic human infections (74). Domestic transmission cycles occur where vectors have become adapted to living in human dwellings and nearby animal enclosures; domestic mammals such as dogs, cats, and guinea pigs play important roles as triatomine blood meal sources and T. cruzi reservoir hosts (69, 124, 131). Some triatomine species can infest both domestic and sylvatic sites and may play a bridging role (192). There are more than 130 triatomine species in the Americas, many of which can be infected by and transmit T. cruzi (169, 311). However, a small number of highly domiciliated vectors are of disproportionate importance in the human epidemiology of disease (Table 2) (311). The domestic environment provides abundant blood meal sources, and poor quality housing with adobe or unfinished brick walls provides crevices and other diurnal hiding places for triatomines (170, 201). Thatch roofs provide an attractive habitat for some species (117). In communities where the disease is endemic, 25 to 100% of houses may be infested, and a house and its immediate surroundings may support large colonies of juvenile and adult bugs (170, 201, 230). In areas of the Amazon where deforestation and human immigration have occurred, tree-dwelling sylvatic triatomine

658

CLIN. MICROBIOL. REV.

BERN ET AL.

TABLE 2. The major triatomine species that colonize the domestic and peridomestic environment and play an important role in the epidemiology of Chagas’ disease in Latin Americaa Vector species

Locations b

c

c

b

Triatoma infestans ..................................................Argentina, Brazil, Chile, Paraguay, southern Peru, Uruguayc Rhodnius prolixus ...................................................Colombia, El Salvador, Guatemala,d Honduras, southern Mexico, Nicaragua, Venezuela Triatoma dimidiata ................................................Belize, Colombia, Costa Rica, Ecuador, El Salvador, Guatemala, Honduras, Mexico, Nicaragua, Panama, northern Peru, Venezuela Panstrongylus megistus ...........................................Argentina, Brazil, Paraguay, Uruguay Triatoma brasiliensis ..............................................Northeastern Brazil a

Data are from reference 311. T. cruzi transmission by T. infestans has been certified as interrupted in 6 provinces of Argentina and 1 department of Paraguay (220). T. cruzi transmission by T. infestans has been certified as interrupted throughout the country (220). d T. cruzi transmission by R. prolixus has been certified as interrupted throughout the country (220). b c

populations have survived and rebounded by adapting to new vertebrate host species (2). These opportunistic vertebrates (opossums and rodents) are competent Chagas’ disease reservoirs and are acclimated to living in close proximity to humans where remnant vegetation is located. The concentration of triatomines and vertebrate reservoirs in the peridomestic realm has lead to increased interactions between sylvatic triatomine species and humans in deforested areas of the Amazon and Panama and to an apparent increase in the incidence of Chagas’ disease in humans (4, 244). Triatomine Distribution in the United States Eleven species of triatomine bugs have been reported from the United States: Triatoma gerstaeckeri, T. incrassata, T. indictiva, T. lecticularia, T. neotomae, T. protracta, T. recurva, T. rubida, T. rubrofasciata, T. sanguisuga, and Paratriatoma hirsuta (Fig. 1 and Table 3). Triatomines are present across the southern half of the country, distributed from the Pacific to Atlantic coasts (Fig. 2). One species (T. rubrofasciata) is found in Hawaii. A high degree of polymorphism has been noted in several species across their geographic ranges, particularly T. protracta, T. rubida, and T. sanguisuga, resulting in proposed subspecies classifications (249, 251, 254, 296). However, due to the recognition of morphological intermediates across some subspecies groups and the absence of supporting data (e.g., paired molecular and morphological studies), these subspecies have not been universally accepted as valid taxonomic groups (110, 169). All U.S. species except T. rubrofasciata and T. sanguisuga

have been collected in Mexico; the distribution of T. sanguisuga likely extends into northeastern Mexico as well (255). A review of the published literature from 1939 to 2010 resulted in reports of wild-caught triatomine bugs from 262 counties in 28 states. The greatest species diversity occurs in the southwest, particularly Texas, Arizona, and New Mexico. More specifically, high species diversity is concentrated in south-central Arizona and southwestern Texas, where up to five species have been recorded in a single county (Fig. 2). T. cruzi-infected specimens have been reported from 10 states, predominantly from counties in the Southwest (Fig. 3A). All species except T. incrassata and P. hirsuta have been found naturally infected with T. cruzi (Fig. 3B to L). County-level maps (Fig. 2 and 3) reflect in part where collection efforts have been focused over the past 70 years. There is no evidence of a temporal or spatial trend in the published reports to suggest any recent migration of species into or within the United States. The county maps do not necessarily reflect triatomine population densities or provide a complete representation of their distributions. Rather, the maps more likely provide an indication of where the bugs have been considered a pest to humans or animals and where field efforts were concentrated as a consequence or where specimens were collected coincidentally by researchers studying other animal systems (i.e., reports based on museum specimens). Collection records are more comprehensive in the southwestern states and Florida, with sparse records in the southeastern states. Early discovery of the association of U.S. triatomine bugs with Neotoma species of woodrats may have aided field research in

FIG. 1. Photographs of U.S. triatomine species, Triatoma and Paratriatoma. The image size relative to the scale bar represents the average length of each species. Photographs for T. incrassata, T. recurva, and P. hirsuta were unavailable. All photographs are by S. Kjos.

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

659

TABLE 3. Geographic location, Trypanosoma cruzi prevalence, human interaction, and sites of collection of Triatoma and Paratriatoma species in the United States Species

State(s)

Total no. tested

No. (%) positive

Human bites/allergic reactions

1,800

1,038 (58)

⫹/⫹

Not reported 4 (33) 144 (51)

⫺/⫺ ⫺/⫺ ⫹/⫺

40 (76) 723 (18)

⫺/⫺ ⫹/⫹

71 (13)

⫹/⫹

1,340

96 (7)

⫹/⫹

2 1031

2 (100) 151 (15)

⫹/⫹ ⫹/⫹

0 (0)

⫹/⫺

T. gerstaeckeri

NM, TX

T. incrassata T. indictiva T. lecticularia

AZ Not reported AZ, NM, TX 12 FL, GA, MO, NM, 282 OK, SC, TN, TX TX 53 AZ, CA, CO, NM, NV, 4,124 TX, UT

T. neotomae T. protracta

T. recurva

AZ

T. rubida

AZ, CA, NM, TX

T. rubrofasciata FL, HI T. sanguisuga AL, AR, FL, GA, IL, IN, KS, KY, LA, MD, MO, MS, NC, NJ, OH, OK, PA, SC, TN, TX, VA P. hirsuta AZ, CA, NV a

565

66

Collection site(s)a

References

B, C, D, H, L, LS, WR

26, 49, 94, 150, 160, 169, 195, 217, 228, 239, 259, 282, 296, 330–332, 341 L 169, 255 H, L, WR 150, 151, 229, 259, 296, 332, 341 D, H, L, T, WR 150, 169, 195, 218, 250, 256, 259, 282, 312, 332, 341 D, WR 49, 85, 94, 150, 282, 296 H, L, R, T, WR 95, 96, 135, 150, 152, 153, 159, 187, 203, 204, 217, 237, 243, 255, 256, 259, 273, 282, 285, 296, 304, 320–322, 326, 327, 329, 330, 332, 335, 336, 339–341 C, H, L, R, WR 95, 96, 152, 237, 255, 256, 296, 321, 325, 329, 330, 332, 335, 336, 339 H, L, WR 95, 96, 150, 152, 153, 156, 237, 256, 259, 273, 282, 296, 321, 324, 329, 330, 332, 335, 336, 341 H, LS, WP 12, 169, 255, 296, 337 D, H, L, LS, T, WP, 27, 41, 49, 54, 77, 90, 94, 116, WR 120, 128, 134, 147, 150, 152, 169, 195, 212, 218, 228, 231, 239, 254, 259, 282, 286, 296, 332, 345, 347 H, L, WR 169, 251, 252, 255, 256, 296, 324, 333, 335, 336

B, bird nest; C, cave; D, dog kennel; H, house; L, lights; LS, livestock pens; R, roadbed; RK, rocks; T, trees; WP, woodpile; WR, woodrat nest.

the southwestern states, because woodrat species in this region build easily identifiable, above-ground dens. The absence of records in some areas of the southeastern United States may reflect a paucity of field studies or published records in those

locations rather than being an indication of true absence of the bug. The detection of T. cruzi-infected wild mammals in many of these areas suggests the presence of the vectors. Additionally, recent efforts to model the geographic distribution of U.S. species based on the land cover, climate, and host composition of known collection sites indicate favorable habitat suitability in many of these unsurveyed or underreported regions (26, 137, 158, 259). Characteristics of each species are summarized in Table 3 and described in detail in the sections that follow.

Description of U.S. Triatomine Species

FIG. 2. Triatomine species diversity in the continental United States and Hawaii by county. States shaded gray have reported at least one species. The states of Kentucky, Maryland, Mississippi, New Jersey, and Pennsylvania have each reported one species but with no locality specified. References are provided in Table 3.

Triatoma gerstaeckeri (Stål). T. gerstaeckeri is one of the most frequently collected and tested species in the United States; 57.7% (1,038/1,800) of tested specimens were found to harbor T. cruzi. T. cruzi-infected specimens have been found in both Texas and New Mexico and in the majority of the counties where testing has been reported (Fig. 3B). Published reports from the 1930s to 1960s describe T. gerstaeckeri as a pest species of humans and livestock; the adult bugs were frequent invaders of rural houses in Texas, and reports of humans being bitten were common (217, 330, 332). Human encounters have been less frequently reported in recent decades (49, 151). Infected T. gerstaeckeri specimens were recently recovered from the residence of a child with acute Chagas’ disease in southern Texas (151). In northeastern Mexico, this species is considered an important Chagas’ disease vector due to its close association

660

BERN ET AL.

CLIN. MICROBIOL. REV.

FIG. 3. Triatomine species geographic distribution by state (gray areas) and county and Trypanosoma cruzi infection status by county in the continental United States and Hawaii. (A) All species; (B) Triatoma gerstaeckeri; (C) T. incrassata; (D) T. indictiva; (E) T. lecticularia; (F) T. neotomae; (G) T. protracta; (H) T. recurva; (I) T. rubida; (J) T. rubrofasciata; (K) T. sanguisuga; (L) Paratriatoma hirsuta. Red, T. cruzi-positive specimens; blue, negative specimens; yellow, no testing reported. References are provided in Table 3.

with human dwellings (184, 288). U.S. T. gerstaeckeri data derive predominantly from Texas, where the bug has been found in a wide variety of habitats. The species was collected from a rock squirrel burrow in a cave in the southeastern corner of New Mexico (341). Triatoma incrassata Usinger. T. incrassata is somewhat similar to T. protracta in size and general appearance of legs and head, but it has a distinctive abdominal margin which is largely yellow on the dorsal surface and entirely yellow on the ventral surface. It has been collected at lights in the two southern Arizona counties of Santa Cruz and Pima (Fig. 3C) (169, 255). The major mammalian hosts and T. cruzi infection prevalence for this species are unknown. Triatoma indictiva Neiva. T. indictiva was considered a subspecies of T. sanguisuga in the past but is currently accorded full species status (110, 169, 296). This species is very similar in appearance to T. sanguisuga, with the exception of the uniformly black pronotum and narrower horizontal markings on the abdominal edge. The distributions of the two species overlap in the central regions of Texas, with T. indictiva continuing

further west to Arizona and T. sanguisuga continuing east to the Atlantic coast (Fig. 3D and K). Reported collection of T. indictiva is much less frequent than that of T. sanguisuga. Additional collection sites for T. indictiva in New Mexico and Arizona were provided in a map by Lent and Wygodzinsky in 1979, but specific location designations were not given (169). Specimens were collected from woodrat nests in New Mexico and at lights in Texas (229, 332). T. indictiva has been found naturally infected with T. cruzi in specimens from Texas (151, 229). Triatoma lecticularia (Stål). T. lecticularia has a geographic distribution similar to that of T. sanguisuga, from the southcentral United States east to the Atlantic coast (Fig. 3E). Its range probably includes Oklahoma, Arkansas, Louisiana, Mississippi, and Alabama based on similarities in ecological characteristics between these states and adjacent areas where it has been reported. Specimens of T. lecticularia from New Mexico have been reported, but specific location information was not provided (254, 296). T. lecticularia had been variously classified as a subspecies of as well as synonymized with T. sanguisuga

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

prior to Usinger’s 1944 reclassification (296). Therefore, early reports of T. lecticularia and T. sanguisuga may be difficult to confirm without reviewing the actual specimens. Ryckman in 1984 contended that reports of T. lecticularia from Arizona and California are erroneous, presumably based on earlier taxonomic confusion and contemporary knowledge of the species distribution (254). T. lecticularia can be distinguished from T. sanguisuga and T. indictiva based on its shorter, domed head and uniform covering of all body surfaces with dark hairs. T. lecticularia has been collected from houses, dog kennels, woodrat nests, and rock squirrel burrows in hollow logs in Texas, from houses in South Carolina, and at lights in Missouri (151, 195, 256, 312, 345). In early reports, this species was described as a nuisance species, commonly found in well-constructed homes of central Texas (218). In 1940, Packchanian conducted experimental inoculation of the gut contents of a T. cruziinfected T. lecticularia bug into the eye of a human subject in order to demonstrate the infectivity of a T. cruzi strain from Texas (216). Localized symptoms, fever, lymphadenopathy, and trypomastigotes visualized on blood films confirmed infection in this individual. The high T. cruzi infection prevalence (144/282; 51%) in T. lecticularia was derived primarily from specimens collected from woodrat nests in Texas (282, 332). Triatoma neotomae Neiva. In the United States, T. neotomae is known only from Texas, primarily the southern tip (Fig. 3F). The inclusion of other states in its range by some authors is most likely an error, as published records of T. neotomae outside Texas or northeastern Mexico could not be found. This species is similar in size to T. protracta but with distinctive yellow markings around the abdominal margin and basal half of wings, a glossy body surface, and a ventrally flattened abdomen. Also like T. protracta, this species is closely associated with Neotoma spp. of woodrats, for which it was named. It has been found almost exclusively in woodrat nests throughout its range, with a single report from a dog kennel in Cameron County, TX (151). The small sample size limits interpretation of this species’ high cumulative T. cruzi infection prevalence (40/53; 76%); however, this is likely related to the high infection levels reported among woodrats in this region (49, 93, 219). Triatoma protracta (Uhler). T. cruzi was first reported in the United States from a T. protracta specimen collected in 1916 in a woodrat nest in San Diego County, CA (155). T. cruzi testing data are most abundant for this species, with an overall prevalence of 17.5% (723/4,124). Infected specimens have been reported from four of seven states across its range: California, Arizona, New Mexico, and Texas (Fig. 3G). T. protracta is closely associated with western woodrat species and is commonly found in nests throughout the bug’s geographic distribution. Large aggregations of T. protracta were reported from roadbeds in southern California in an area where woodrat nests were removed as a consequence of highway construction (340). Attracted by lights, the displaced bugs frequently entered houses in the area and became a source of annoyance for residents. T. protracta has also been reported as frequently entering houses in other areas of California, New Mexico, and Arizona (187, 273, 304, 332, 336). First reported as a pest of humans in Yosemite Valley, CA, in the 1860s, T. protracta continues to be an important cause of severe allergic reactions in humans who are bitten (152, 198). This species was impli-

661

cated in a human case of Chagas’ disease in north-central California (203). Triatoma recurva (Stål). T. recurva naturally infected with T. cruzi has been found in the southern half of Arizona (Fig. 3H). A single report of T. recurva collected in western Texas has not been confirmed or replicated (138, 151). Early reports describe T. recurva as a pest of humans, primarily in the Alvardo Mine area of Yavapai County, AZ, where it was a common invader of houses and tents of mining employees (332, 336). Recent reports describe home invasions and hypersensitivity reactions due to bites that occurred in and around houses in Pima and Cochise Counties, AZ (152, 237). Although the species has been collected occasionally from woodrat nests, the woodrat is not considered the primary host of T. recurva (96, 255, 321). The preferred host for this species is unknown, but it has been observed in association with rodents, particularly rock squirrels, and feeds on reptiles and guinea pigs in laboratory settings (96, 255, 324, 334, 336). T. recurva is the largest of the U.S. species (average length, 29 mm) and has relatively hairless body surfaces, including the first two segments of the mouthparts. It is brown to black in appearance, with slender, long legs and head and an orange-yellow abdominal margin. Its body size, head and leg characteristics, and uniformly colored pronotum distinguish this species from others in its range. Triatoma rubida (Uhler). In the United States, T. rubida has been found from western Texas to southern California; T. cruzi-positive specimens have been reported from Arizona and Texas (Fig. 3I). The cumulative infection prevalence in the published literature is low (96/1,340; 7.2%). However, in a recent study, the gut contents of 65 (41%) of 158 T. rubida specimens collected in and around houses in Pima County, AZ, yielded positive results by T. cruzi PCR (237). Despite the presence of nymphal stages inside houses in this study, the authors remarked that the numbers were too low to conclude that colonization was established. In contrast, a study from Sonora, Mexico, reported that 68% of houses were colonized by T. rubida, suggesting that this species was domesticated in that region (221). Both the U.S. and Mexican study areas had experienced disruption of previously undisturbed environments considered suitable habitats for both triatomine and T. cruzi vertebrate hosts. Human bite encounters, including hypersensitivity reactions due to T. rubida, continue to be a public health issue in Arizona (152, 226, 237). This species has been frequently collected from woodrat nests throughout its range (96, 256, 321, 332, 336). It can be distinguished morphologically from other species in its range by the first antennal segment, which reaches or surpasses the tip of the head. Triatoma rubrofasciata (DeGeer). Described in 1733, T. rubrofasciata was the first species classified in the Triatominae subfamily and is the current type species for the Triatoma genus (270). It is the only triatomine species found in both the Eastern and Western Hemispheres and is frequently found in port cities in close association with the roof rat (Rattus rattus) (255). Molecular and morphometric data support the hypothesis that Old World triatomine species derive from T. rubrofasciata carried from North America with rats on sailing ships during the colonial period (136, 223, 270). In the United States, this species has been collected from houses in Florida and Hawaii and in chicken and pigeon coops and cat houses in

662

BERN ET AL.

Hawaii. Specimens have been reported from Jacksonville, FL, and Honolulu, HI (Fig. 3J) (296, 337). Wood (in 1946) reported 2 specimens collected from Honolulu to be infected with T. cruzi based on morphological and motility characteristics (337). Allergic reactions to T. rubrofasciata bites have been reported in humans from Hawaii (12). Triatoma sanguisuga (Leconte). T. sanguisuga is one of the most widely distributed species in the United States, with its range spanning from Texas and Oklahoma eastward to the Atlantic coast (Fig. 3K). This species has been reported in Pennsylvania, New Jersey, Maryland, and Kentucky, but without specific location data (169, 254, 296). Although published records are lacking, its range probably includes West Virginia. Reports of T. sanguisuga from states west of Texas were likely mistaken due to taxonomic reclassification (see “Triatoma indictiva Neiva” above). In every state where testing has been conducted, T. cruzi-infected T. sanguisuga has been found, including Texas, Oklahoma, Louisiana, Alabama, Tennessee, Georgia, and Florida. It has been collected from diverse natural settings across its range, in association with many different vertebrate hosts, including woodrats, cottonrats, armadillos, raccoons, opossums, frogs, dogs, chickens, horses, and humans (120, 150, 212, 215, 332, 348). Human annoyance and allergic reactions to T. sanguisuga bites were reported as early as the mid-1800s in Georgia, Kansas, Oklahoma, and Florida and recently in Louisiana (116, 147, 152, 161, 215). This species was found inside the residences of human Chagas’ disease patients in Tennessee and Louisiana and in the vicinity of the home of a T. cruzi-seropositive blood donor in Mississippi (54, 90, 134). Paratriatoma hirsuta Barber. P. hirsuta is known from the western United States, collected from arid regions of California, Nevada, and Arizona (Fig. 3L). Although it has been demonstrated to be a competent vector of T. cruzi in experimental settings, a naturally infected specimen has yet to be reported (321). It has been most frequently collected from woodrat nests in its range but has also been found in houses and other human dwellings in Yavapai County, AZ, and Riverside County, CA, and at lights in Palm Springs, CA (251, 296, 336). Ryckman (in 1981) described this species as having important public health significance due to allergic reactions caused by its bite (252). This is one of the smallest U.S. triatomine species (average length, 13 mm) and can be distinguished from T. protracta, which is similar in size and geographic distribution, by a pervasive covering of dark hairs on all body surfaces. Human-Vector Interactions and T. cruzi Transmission Potential in the United States Eight of the 11 species have been associated with human bites, and seven have been implicated in allergic reactions (Table 3). Allergic reactions occur in response to antigens delivered in the vector saliva during blood feeding and are unrelated to the T. cruzi infection status of the bug. Most allergic reactions are localized at the bite site, characterized by a large welt and intense itching (315). Severe reactions are generally systemic and may involve angiodema, urticaria, difficulty breathing, nausea, diarrhea, and/or anaphylaxis (152, 226). Although allergic reactions to triatomine bites have been

CLIN. MICROBIOL. REV.

reported from states throughout the southern United States, the incidence is highest in the southwestern states, with T. protracta and T. rubida most frequently implicated (106, 152, 204, 226, 237). The most common scenario involves invasion of an adult bug into a human dwelling, where it bites a sleeping individual. Contemporary encounters between humans and triatomine bugs in the United States are often associated with destruction or invasion of vertebrate host habitats, compromised housing structures, or both. Disruption of host burrows (as described above for T. protracta) provokes the bugs to seek new refuges, and their innate attraction to lights often leads them to nearby human dwellings. Most triatomine species show flexibility in host and habitat requirements, which allows them to adapt to changing environments. A host preference for some species has been difficult to establish due to association with multiple vertebrate habitats and the ability of the insects to mature and reproduce successfully on multiple host species in laboratory settings. Although mammals are the only vertebrate reservoirs for T. cruzi, many triatomine species utilize other animal groups as blood hosts, including reptiles and amphibians (T. gerstaeckeri, T. protracta, T. recurva, T. rubida, and T. sanguisuga) and birds (T. gerstaeckeri and T. sanguisuga) (169, 228, 253, 338). A recent blood meal analysis study of Texas field specimens provides evidence of a broad host range for T. gerstaeckeri and T. sanguisuga. The DNAs from nine vertebrate species (woodrat, dog, cat, cow, pig, raccoon, skunk, armadillo, and human) were detected in T. gerstaeckeri gut specimens, and DNAs from three species (dog, avian, and human) were detected in T. sanguisuga gut specimens (149). Because vector colonization of houses in the United States is rare, the risk of vector-borne transmission to humans is considered to be quite low. With the exception of the 2006 Louisiana case in which the residence was found to harbor triatomine colonies, vector-borne transmission to humans in the United States has been attributed to adult bugs invading houses (90, 134, 203). Expansion of human settlements into environments that support an active sylvatic disease cycle could result in an increase in adult invaders and, potentially, colonization events. Colonization of houses by triatomines is an important factor in vector-borne transmission because it increases the probability of encounters between humans and potentially infected vectors. In addition to adaptability to domestic structures, triatomine feeding and defecation behaviors are important risk factors for vector-borne transmission and vary across species. The timing and placement of defecation after feeding greatly influence the risk of transmission via fecal contamination of the host bite site or other exposed tissues. A small number of studies have reported on these characteristics in U.S. species. In 1951 Wood reported the following average postfeeding defecation times (minutes) for the adults of four U.S. species: T. protracta, 30.6 (n ⫽ 10); T. rubida, 1.6 (n ⫽ 5); T. recurva, 75.7 (n ⫽ 3); and P. hirusta, 35.0 (n ⫽ 2) (327). In a similar study in 2007 using both nymphs and adults of three Mexican species (also present in the United States), Martinez-Ibarra et al. reported the following results: T. protracta, 6.7 (n ⫽ 475); T. lecticularia, 8.3 (n ⫽ 368); and T. gerstaeckeri, 11.5 (n ⫽ 733) (183). Likewise, Zeledon et al. (1970) reported the following results for nymphs and adults of three Latin American species: R. prolixus, 3.2

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

(n ⫽ 210); T. infestans, 3.5 (n ⫽ 210); and T. dimidiata, 11.3 (n ⫽ 210) (352). In 1970 Pippin reported the proportion of bugs defecating within 2 min postfeeding for adults of two U.S. and one Latin American species: R. prolixus, 74.6% (n ⫽ 169); T. gerstaeckeri 19.4% (n ⫽ 160); and T. sanguisuga 16.9% (n ⫽ 136) (228). Similarly, in 2009 Klotz et al. reported the proportion of bugs defecating before or directly after feeding for adults of two U.S. species: T. rubida, 45% (n ⫽ 40); and T. protracta, 19.4% (n ⫽ 31) (153). In that study, it was noted that none of the bugs of either species defecated on the host during the experiment. Although direct comparisons across studies is problematic due to variation in methods and conditions (e.g., temperature, blood host, and feeding apparatus), it appears that U.S. species in general exhibit greater postfeeding defecation delays than important Latin American vector species. Delayed defecation and a low frequency of domestic colonization contribute to a low probability of autochthonous U.S. human infection due to vector-borne transmission, which is the primary route of infection in areas of hyperendemicity in Latin America.

ANIMAL RESERVOIRS OF TRYPANOSOMA CRUZI Background Concurrent with the demonstration of T. cruzi in the first recognized human patient, Carlos Chagas observed the parasite in the blood of a domestic cat in the same household (62). Subsequently, Chagas went on to demonstrate T. cruzi in armadillos and primates, confirming the role of wildlife as reservoirs (63, 64). To date, over 100 mammalian species have been reported as natural hosts for T. cruzi, and all mammals are considered to be susceptible to infection. Birds are refractory to infection due to complement-mediated lysis and macrophage-induced killing of the parasites (146, 188). Although T. cruzi has a wide host range, the epidemiologically important reservoirs vary by geographic region due to the biology and ecology of the mammals and vectors and how these interactions translate to risk of human exposure. Opossums and armadillos are important reservoirs throughout the Americas, a finding consistent with genetic data suggesting that these two groups are the ancestral hosts for the two major ancestral lineages of T. cruzi (112, 349). Transmission routes for wildlife and domestic animal species are similar to those for humans. Sylvatic animals become infected during feeding activity of vectors present in their burrows, dens, or temporary shelters. As in humans, infection occurs when bug feces containing the parasites enters a wound or mucous membrane. In addition, the insectivorous behavior of many animals (e.g., woodrats) increases the likelihood of infection via the ingestion of infected bugs (78, 241, 250). Transplacental transmission has been documented in laboratory mice and rats (15, 81, 127). Although not proven to occur in wildlife, this route likely contributes to maintenance of the parasite in the sylvatic cycle as well. Ingestion of infected meat was once considered a possible route of transmission, but a recent study with raccoons suggests that this is probably uncommon (241).

663

Wildlife Reservoirs of T. cruzi in the United States In the United States, natural T. cruzi infection was first reported in the big-eared woodrat Neotoma macrotis (syn. N. fuscepes macrotis) in California (316). In the 1940s, natural infections were reported from house mice, southern plains woodrats (N. micropus), nine-banded armadillos (Dasypus novemcinctus), and Virginia opossums (Didelphis virginiana) in Texas and from brush mice (Peromyscus boylii rowleyi) and woodrats (N. albigula) in Arizona (219, 321). Early experimental infection trials with parasites from these hosts and Triatoma spp. indicated that laboratory rats, laboratory mice, guinea pigs, domestic dogs, rhesus macaques, opossums (D. virginiana), six species of Peromyscus, and four species of woodrats were susceptible (77, 154, 215, 217, 316, 321). In addition, an isolate from a naturally infected Triatoma species from Texas was shown to be infectious to a human (216). Subsequent surveys in the 1950s and 1960s documented infections in raccoons (Procyon lotor), Virginia opossums, striped skunks (Mephitis mephitis), and gray foxes (Urocyon cinereoargenteus) in the southeastern United States (185, 212, 305). Currently, at least 24 species are recognized as natural wildlife hosts for T. cruzi in the United States (Table 4). Reported T. cruzi infection rates vary widely by host species and geographic area. However, the observed variation may be due in part to the use of different diagnostic assays with very different sensitivities. As in humans, the majority of infected animals are in the chronic phase of the infection; therefore, serological testing is more sensitive than methods that rely on detection of parasites (346). However, unlike serological tests, visualization of the parasites allows the examiner to distinguish T. cruzi from other Trypanosoma species (e.g., T. neotomae, T. kansasensis, T. peromysci, and T. lewisi-like sp.) reported from rodents based on morphology (186, 207, 294, 317, 328; M. J. Yabsley, unpublished data). If serology is used for screening, infections should be confirmed with a T. cruzi-specific assay. Some PCR assays will amplify other Trypanosoma and/or Leishmania species, and more specific methods may be necessary to confirm the infection as T. cruzi. The primary reservoirs and transmission dynamics of T. cruzi differ between the eastern and western regions of the United States. The greatest vector diversity and density occur in the western United States (Fig. 2), where many triatomine species live in the nests of woodrats. In this region, woodrats are the most common reservoir; however, infection has also been demonstrated in other rodents, raccoons, skunks, and coyotes (Table 4; Fig. 4A and B). Rodents other than woodrats utilize habitats similar to those of woodrats (old woodrat nests, small caves, and holes in rock walls) where triatomines are found, while coyotes, raccoons, skunks, and opossums likely become infected when bugs feed on them in their dens or through ingestion of bugs. In the eastern United States, the prevalence of T. cruzi is highest in raccoons, opossums, armadillos, and skunks (Table 4; Fig. 4A and B). There are several woodrat species in the eastern United States, but densities are much lower than for woodrat species in the western United States, and nests are less evident because they utilize burrows instead of large above-ground constructed nests. Little is known about the prevalence of T. cruzi in eastern woodrat species. To date, only one survey for T. cruzi has been conducted, and none of 23

664

BERN ET AL.

CLIN. MICROBIOL. REV. TABLE 4. Hosts of Trypanosoma cruzi in the United Statesa

Species

Raccoon (Procyon lotor)

State(s)

Total no. tested

AL AZ FL FL FL FL, GA GA GA GA GA GA, SC KY

35 5 184 33 70 608 54 30 510 10 221 44

MD MD MO NC OK TN TN TX TX TX

No. (%) positive

Assay type (sample or specific assay)

Reference(s)

472 NKb 109 20 8 3 706 25 9 19

5 (14) 1 (20) 4 (2) 4 (12) 38 (54) 9 (1.5) 12 (22) 13 (43) 168 (33) 5 (50) 104 (47) 17 (39) 19 (43) 2 (0.4) 5 74 (68) 3 (15) 5 (63) 2 (66) 206 (29) 6 (24) 0 4 (21)

Culture (heart and blood) Serology (IFA) Blood smear Culture (blood) Serology (IFA) Culture (kidney) Culture (blood) Culture (blood) Serology (IFA) Culture (blood) Serology (IFA) Culture (blood) Serology (IFA) Culture (heart) Culture (blood) Serology (IFA) Culture (blood) Culture (blood) Culture (blood) Serology (IFA) Culture (blood) Serology (indirect hemagglutination) Culture (blood)

VA

464

153 (33)

Serology (IFA)

130 305 46 144 141 134 179 262 49 M. Yabsley et al., unpublished 129

Ringtail (Bassariscus astutus)

AZ

1

Serology (IFA)

46

Opossum (Didelphis virginiana)

AL FL GA GA GA GA, FL KY

126 27 39 421 29 552 48

212 46 231 46 222 185 118

MD NC OK LA TX TX VA

219 12 10 48 8 391 6

Culture (blood and heart) Serology (IFA) Culture (blood) Serology (IFA) PCR (liver) Culture (kidney) Culture (blood) Serology (IFA) Culture (heart) Culture (blood) Culture (blood) Culture (blood) Culture (blood) Blood smear Serology (IFA)

LA LA

98 80

19 348

TX

15

Culture (blood) Culture (blood) Serology (direct agglutination) Culture (blood)

Nine-banded armadillo (Dasypus novemcinctus)

1 (100) 17 (14) 14 (52) 6 (15) 118 (28) 3 (10) 88 (16) 0 (0) 6 (13) 0 (0) 1 (8) 0 16 (33) 5 (63) 63 (16) 1 (17) 1 (1) 23 (29) 30 (38) 1 (7)

212 46 262, 284 262 46 185 231 224 46 262 345 118

130 144 141 19 219 94 46

219

Striped skunk (Mephitis mephitis)

AZ CA GA, FL GA TX

34 1 306 1 3

3 (9) 1 (100) 3 (1) 1 (100) 2 (67)

Serology (IFA) Serology and histology Culture (kidney) Serology (IFA) Culture (blood)

46 248 185 46 Yabsley et al., unpublished

Gray fox (Urocyon cinereoargenteus)

GA, FL GA SC

118 21 26

2 (2) 0 2 (8)

Culture (kidney) Serology (IFA) Serology (IFA)

185 46 245

Bobcat (Felis rufus)

GA

62

2 (3)

Serology (IFA)

46

American badger (Taxidea taxus)

TX

8

2 (25)

Serology (indirect hemagglutination)

49

Coyote (Canis rufus)

GA TX

23 134

1 (4) 19 (14)

Serology (IFA) Serology (IFA)

46 119 Continued on following page

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

665

TABLE 4—Continued Total no. tested

No. (%) positive

VA

26

1 (4)

Serology (IFA)

46

Feral swine (Sus scrofa)

GA

110

0

Serology (IFA)

46

Southern plains woodrat (Neotoma micropus)

TX TX TX NM

30 100 159 NK

Culture (blood) Culture (blood) PCR (liver) Xenodiagnosis

49 219 227 341

White-throated woodrat (Neotoma albigula)

AZ NM

NK NK

2 1

NK Xenodiagnosis

328 341

Big-eared woodrat (Neotoma macrotis ⫽ N. fuscipes subsp. macrotis) Brush mouse (Peromyscus boylii rowleyi) Gilbert white-footed mouse (Peromyscus truei gilberti) Pinon mouse (Peromyscus truei montipinoris) Western harvest mouse (Reithrodontomys megalotis) Hispid pocket mouse (Perognathus hispidus) House mouse (Mus musculus) Mexican spiny pocket mouse (Liomys irrorattus) Grasshopper mouse (Onychomys leucogaster) CA ground squirrel (Spermophilus beecheyi) Mexican ground squirrel (Spermophilus mexicanus) Whitetail antelope squirrel (Ammospermophilus leucurus) Hispid cotton rat (Sigmodon hispidus)

CA

99

9 (9)

Xenodiagnosis and blood smear

318, 319, 328, 339

AZ

NK

1

NK

328

CA

NK

2

NK

328

CA

NK

11

Xenodiagnosis

339

CA

NK

1

Xenodiagnosis

323

TX

25

4 (16)

Culture (blood)

49

TX TX

2 11

1 (5) 1 (9)

Culture (blood) Culture (blood)

219 49

TX

9

1 (11)

Culture (blood)

49

CA

19

2 (11)

Culture (blood)

203

TX

1

1 (100)

Culture (blood)

NM

NK

3

Xenodiagnosis

Yabsley et al., unpublished 339, 341

TX

1

1 (100)

Culture (blood)

Species

a b

State(s)

5 (17) 31 (31) 42 (26) 1

Assay type (sample or specific assay)

Reference(s)

Yabsley et al., unpublished

Only selected negative results are shown if large numbers of a particular species were examined. NK, not known.

Neotoma floridana animals in Kansas were positive (294). Reports of wildlife infections are shown at the county level (when possible) in Fig. 4A and B. Domestic and Exotic Animal Infections in the United States In addition to indigenous wildlife reservoirs, domestic and exotic animals can become infected if they are present in an enzootic area and come in contact with infected bugs. Transmission routes are similar to those for wildlife, with ingestion of bugs likely being an important route. Canine Chagas’ disease. In Central and South America, domestic dogs are important reservoirs in the domestic cycle and can be used as sentinels for local transmission (123). A similar cycle has been recognized in the United States, but the importance of domestic dogs as T. cruzi infection reservoirs is not as well understood (26). T. cruzi infection in domestic dogs has been reported widely throughout the southern United States since 1972 (Fig. 4C) (18, 20, 23, 41, 105, 134, 150, 189, 205, 206, 239, 274, 287, 298, 312). Infection has been documented in at least 48 different breeds in the United States, with

the sporting and working breeds accounting for the majority of cases, presumably due to greater exposure to infected vectors and mammalian tissues (150, 246). As in humans, transplacental transmission is also an important mode of transmission in dogs (23, 58). Domestic dogs can develop both acute and chronic disease similar to that in humans. Acute illness, particularly mortality, has been reported more frequently in very young dogs (⬍1 year old) and generally involves myocarditis and cardiac arrhythmias (150). Dogs that survive infection at a very young age or acquire infection as adults generally experience a chronic course of disease that may progress to significant cardiac dysfunction, typically involving cardiac dilatation, electrocardiogram (ECG) abnormalities, and clinical signs related to right-sided or bilateral cardiac failure (21, 22). In a recent seroprevalence study in Tennessee, older dogs (ages 6 to 10 years) were more likely to be infected (246), which is similar to results of studies in Latin America that reported increasing seropositivity with increasing age (97, 122). Dogs with clinically apparent infections are managed with appropriate supportive therapy. Chemotherapeutic agents developed for treatment of human Chagas’ disease (benznidazole and

666

BERN ET AL.

CLIN. MICROBIOL. REV.

inoculum from exposure to or ingestion of a large number of infected bugs but may also reflect variation in susceptibility of animal species to clinical Chagas’ disease. Mortality due to locally acquired T. cruzi infection has occurred in groups of captive animals in the United States, including baboons (Papio hamadryas), rhesus macaques, crab-eating macaques (M. fascicularis), Celebes black macaques (M. nigra), sugar gliders (Petaurus breviceps), and a hedgehog (Atelerix albiventris) (8, 115, 145, 213, 313). Asymptomatic T. cruzi infection has been reported in lion-tailed macaques (M. silenus), pigtailed macaques (M. nemestrina), rhesus macaques, baboons, ring-tailed lemurs (Lemur catta), and black and white ruffed lemurs (Varecia variegata) in the United States (11, 128, 145, 232, 264). MOLECULAR EPIDEMIOLOGY OF T. CRUZI General Molecular Epidemiology

FIG. 4. Reports of natural Trypanosoma cruzi infection in U.S. mammals. (A) Raccoons, Virginia opossums, and ringtails; infection of opossums has been reported in Virginia, but no locality was specified. (B) Rodents and mesomammals. An additional report of infected coyotes was published from Virginia, but no locality was specified. (C) Domestic canines. In some states (California, Georgia, Tennessee, and Virginia, shown in dark gray), additional canine clinical cases were reported, but no locality was specified. References are provided in Table 4 for panels A and B and in the text for panel C.

nifurtimox) have shown some efficacy in dogs (121, 125), but they are not currently approved for veterinary use in the United States. Primates and other exotic animals. Any mammals kept in areas where bugs may enter are at risk of acquiring T. cruzi infection. Because U.S. animal use guidelines require that nonhuman primates be housed in facilities with access to the outdoors, they may be at particular risk of acquiring T. cruzi infection. Exotic animals that acquire T. cruzi infection may be asymptomatic or may develop symptomatic, even lethal, clinical disease. Severe disease may be due to a large parasite

T. cruzi is a genetically heterogeneous species that also has wide variability in biological and biochemical characteristics (51, 174, 191, 192). The most common historical classification divided T. cruzi into two major groups, TcI and TcII; TcII was further divided into five subgroups (also called discrete typing units) designated TcIIa to TcIIe (51, 193, 309). Recently, a consensus was reached that the six major recognized lineages will be renamed TcI to TcVI; compared to the earlier system, TcI remained TcI, TcIIb became TcII, TcIIc became TcIII, TcIIa became TcIV, TcIId became TcV, and TcIIe became TcVI (353). For the purposes of this review, data from earlier studies that genotyped isolates as TcII (without a to d subtyping) will be referred to as “historic TcII” to differentiate these types from the current TcII, which is equivalent to the historic TcIIb lineage. The TcI and TcII lineages are considered ancestral, whereas the TcV and TcVI lineages are the products of at least two hybridization events (309, 353). The origins of TcIII and TcIV are as yet unresolved (353). Whereas some investigators consider TcIII to represent a third ancestral strain (80), others consider it to be the result of hybridization between TcI and TcII (309, 310). TcI and TcII to TcVI are estimated to have diverged between 88 and 37 million years ago (43, 175). Currently, T. cruzi genotypes are classified based on size polymorphism or sequence analysis of several gene loci, including the miniexon gene, the intergenic region of the miniexon gene, the 18S rRNA gene, the 24S␣ rRNA gene, internal transcribed spacer regions, and numerous housekeeping genes (171, 310). The TcI lineage is found throughout the Americas in both domestic and sylvatic cycles and is believed to have evolved with arboreal Didelphimorpha (opossums) and vectors in the triatomine tribe Rhodniini (112). In all parts of the Americas, Didelphis spp. are common reservoirs for this lineage, although natural infection with TcI has been reported in a wide range of mammals. TcI is the only lineage reported from humans in North and Central America and the predominant lineage reported in human Chagas’ disease in areas of South America north of the Amazon Basin (40, 139, 239, 258). Although TcI has long been recognized as genetically diverse, subtyping has not been widely conducted until very recently, and no generally accepted typing system or nomenclature currently exists. Additionally, many isolates have been

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

667

TABLE 5. Genotypes of U.S. T. cruzi isolates Species

No.

State(s)

Genotype(s) (no.)

Reference(s)

Human Opossum Raccoon Ring-tailed lemur Rhesus macaque Nine-banded armadillo Striped skunk Domestic dog Triatoma spp.

5 15 79 3 2 3 1 7 8

CA, TX, LA GA, FL, LA, AL GA, FL, TN, MD, LA, KY GA GA LA, GA GA TN, OK, SC, CA, unknown GA, FL, TX

TcI (5) TcI (15) TcI (2), TcIV (74), mixed (2) TcIV (3) TcI (1), mixed (1) TcI (2), TcIV (1) TcIV (1) TcIV (6), mixed (1) TcI (6), TcIV (1) mixed (1)

239 68, 239 45, 68, 239 239 239 239 239 44, 45, 239 17, 68, 175, 239

examined only by sequencing of a single locus. Haplotypes were first recognized following sequence analysis of the intergenic regions of the miniexon genes of 12 isolates from Columbia (132). Based on single-nucleotide polymorphisms, insertions, and deletions, four haplotypes, TcIa to TcId, were proposed. Haplotypes TcIa and TcIc were associated with humans and domiciliated vectors. Haplotypes TcIb and TcId were found in specimens from one human, opossums, and sylvatic vectors; TcId was found exclusively in sylvatic samples (132). Interestingly, phylogenetic analysis of the same gene region of 20 TcI strains from the United States, Mexico, Bolivia, Brazil, Columbia, and Argentina showed that Didelphis sp. isolates grouped separately from other isolates (210). A fifth haplotype (TcIe) was recently detected in a human and a sylvatic vector (Mepraia spinolai) from Chile and in one domestic vector (T. infestans) from Argentina (76). Multilocus microsatellite profiling of 135 TcI isolates provided better discrimination and increased levels of variability among TcI sylvatic strains (173). However, in contrast to previous studies in which opossums were found to be infected with a particular haplotype (210), no host association was noted. The authors suggest that the ecological niche might be more important for parasite evolution and diversification than reservoir host species (173). Wider use of multilocus typing methods may provide further insight into TcI genetic diversity in the future. Lineage TcIII (historical TcIIc) is believed to have evolved with terrestrial burrowing edentates, specifically armadillos, and bugs in the triatomine tribe Triatomini (112, 349). Edentates and marsupials were the first mammal inhabitants of South America (⬃65 million years ago), whereas rodents, primates, and bats arrived in South America ⬃25 million years ago (112). Upon arrival, these mammals became hosts to the various lineages of T. cruzi. TcIII is found throughout South America and is rare in domestic cycles but common in sylvatic cycles (172, 349). The primary hosts for TcIII are several different species of armadillos, primarily D. novemcinctus. Additionally, TcIII has been reported from limited numbers of a terrestrial marsupial (Monodelphis domestica), rodents, and skunks (349). TcII, TcV, and TcVI (historic IIb, IId, and IIe, respectively) are the lineages most commonly reported in human Chagas’ disease in southern South America (193). All three lineages are closely associated with the domestic transmission cycle and the domestic vector Triatoma infestans. TcV and TcVI have been reported in cardiomyopathy and intestinal megasyndromes in the Southern Cone (Argentina, Chile, Bolivia, and Brazil) (57, 193). In contrast, intestinal Chagas’ disease is rare

in northern South America, Central America, and Mexico, where TcI is the predominant lineage (191). TcV is the lineage reported most frequently in infants with congenital infection, although this may simply reflect its predominance in Bolivia and Argentina, where these studies have been conducted (72, 303). The two hybrid lineages, TcV and TcVI, are hypothesized to have evolved in armadillos (181, 349). A single isolation of TcII (historic TcIIb) was reported from Euphractus sexcinctus (six-banded armadillo) in Paraguay, but its original mammalian host has not been established (193, 349). Currently, the TcIV (historic IIa) lineage is poorly understood. Studies of several gene targets indicate that TcIV strains from North and South America are genetically distinct and group separately in phylogenetic analyses (181; D. M. Roellig and M. J. Yabsley, unpublished data). In South America, TcIV is found in a wide range of mammals, including primates, rodents, armadillos, and terrestrial marsupials (349). In North America, the raccoon is the principal host for TcIV; infections have been reported in domestic dogs, striped skunks, armadillos, and primates (239). Interestingly, two terrestrial marsupial genera (Philander and Monodelphis) can harbor both TcI and several other genotypes, whereas only TcI has been reported from the arboreal genus Didelphis (163, 225, 240). The genus Philander also displays a more severe inflammatory response to T. cruzi (162, 163). Experimental inoculation of Monodelphis domestica with TcI, TcII, and TcVI strains resulted in infections, but a North American TcIV isolate failed to establish an infection (242). Collectively, these data suggest that the marsupial genera diverged before the establishment of host relationships with T. cruzi and that utilization of different ecological niches resulted in distinct T. cruzi lineage transmission patterns (163). T. cruzi Genotypes in the United States In the United States, only two genotypes (TcI and TcIV) have been reported from mammals and vectors (Table 5). Consistent with the findings in South American studies of Didelphis, TcI is the only genotype reported from D. virginiana, the Virginia opossum (17, 68, 239). In raccoons, TcIV predominates, but TcI has been detected in a small number of specimens. Both TcI and TcIV have been reported from ninebanded armadillos, domestic dogs, and rhesus macaques (68, 239). Lineage TcIV has been reported from a limited number of ring-tailed lemurs and a striped skunk (239). Although the majority of isolates from placental mammals in the United States have been TcIV, all five typed isolates from human

668

BERN ET AL.

autochthonous infection were TcI (239). Both TcI and TcIV have been reported from Triatoma spp. from Georgia, Florida, and Texas. CLINICAL ASPECTS OF CHAGAS’ DISEASE Acute T. cruzi Infection The incubation period following vector-borne T. cruzi exposure is 1 to 2 weeks, after which the acute phase begins (234). The acute phase lasts 8 to 12 weeks and is characterized by circulating trypomastigotes detectable by microscopy of fresh blood or buffy coat smears. Most patients are asymptomatic or have mild, nonspecific symptoms such as fever and therefore do not come to clinical attention during the acute phase. In some patients, acute infection is associated with inflammation and swelling at the site of inoculation, known as a chagoma. Chagomas typically occur on the face or extremities; parasites may be demonstrated in the lesion. Inoculation via the conjunctiva leads to the characteristic unilateral swelling of the upper and lower eyelids known as the Roman ˜a sign (234). Severe acute disease occurs in fewer than 1% of patients; manifestations include acute myocarditis, pericardial effusion, and/or meningoencephalitis (3, 177). Children younger than 2 years appear to be at higher risk of severe manifestations than older individuals. Severe acute Chagas’ disease carries a substantial risk of mortality. Orally transmitted T. cruzi infection appears to be associated with more severe acute morbidity and higher mortality than vector-borne infection (28, 271). For example, 75% of 103 infected individuals in the Caracas outbreak were symptomatic, 59% had ECG abnormalities, 20% were hospitalized, and there was one death from acute myocarditis (82). Recent laboratory data suggest that parasite contact with host gastric acid may render trypomastigotes more invasive through changes in parasite surface glycoproteins and that this interaction may underlie the increased clinical severity seen in orally acquired Chagas’ disease (75, 350). Congenital T. cruzi Infection Most infected newborns are asymptomatic or have subtle findings, but a minority present with severe life-threatening disease (32, 289). The manifestations of symptomatic congenital Chagas’ disease can include low birth weight, prematurity, low Apgar scores, hepatosplenomegaly, anemia, and thrombocytopenia (35, 36, 177, 289). Severely affected neonates may have meningoencephalitis, gastrointestinal megasyndromes, anasarca, pneumonitis, and/or respiratory distress (35–37, 289). Mortality among infected infants is significantly higher than in uninfected infants, ranging from ⬍5% to 20% in published studies (34, 289). However, even severe congenital Chagas’ disease may not be recognized because signs are often nonspecific or because the diagnosis is not considered (289). Chronic T. cruzi Infection Eight to 12 weeks after infection, parasitemia levels become undetectable by microscopy, and in the absence of effective etiological treatment, the individual passes into the chronic

CLIN. MICROBIOL. REV.

phase of T. cruzi infection. Despite the absence of microscopically detectable parasites in the peripheral blood, persons with chronic T. cruzi infection maintain the potential to transmit the parasite to the vector and directly to other humans through blood components, through organ donation, and congenitally (177, 311). Indeterminate form of chronic T. cruzi infection. Persons with chronic T. cruzi infection but without signs or symptoms of Chagas’ disease are considered to have the indeterminate form. The strict definition of the indeterminate form requires positive anti-T. cruzi serology, with no symptoms or physical examination abnormalities, normal 12-lead ECG, and normal radiological examination of the chest, esophagus, and colon (194). Current baseline evaluation guidelines in the United States recommend only a history, physical examination, and ECG (30). Further cardiac evaluation is recommended only if cardiac signs or symptoms are present, and barium studies are recommended only in patients with gastrointestinal symptoms (30). An estimated 20 to 30% of people who initially have the indeterminate form of Chagas’ disease progress over a period of years to decades to clinically evident cardiac and/or gastrointestinal disease (234). Cardiac Chagas’ disease. Chagas’ cardiomyopathy is characterized by a chronic inflammatory process that involves all chambers, damage to the conduction system, and often an apical aneurysm. The pathogenesis is hypothesized to involve parasite persistence in cardiac tissue and immune-mediated myocardial injury (182). The earliest manifestations are usually conduction system abnormalities, most frequently right-bundle branch block or left anterior fascicular block, and segmental left ventricular wall motion abnormalities (178). Later manifestations include complex ventricular extrasystoles and nonsustained and sustained ventricular tachycardia, sinus node dysfunction that may lead to severe bradycardia, high-degree heart block, apical aneurysm usually in the left ventricle, thromboembolic phenomena due to thrombus formation in the dilated left ventricle or aneurysm, and progressive dilated cardiomyopathy with congestive heart failure (233). These abnormalities lead to palpitations, presyncope, syncope, and a high risk of sudden death (235, 236). Digestive Chagas’ disease. Gastrointestinal involvement is less common than Chagas’ heart disease. This form is seen predominantly in patients infected in the countries of the Southern Cone (Argentina, Bolivia, Chile, Paraguay, Southern Peru, Uruguay, and parts of Brazil) and is rare in northern South America, Central America, and Mexico. This geographical pattern is thought to be linked to differences in the predominant T. cruzi genotypes (51, 192). Gastrointestinal Chagas’ disease usually affects the esophagus and/or colon, resulting from damage to intramural neurons (83, 84, 199). The effects on the esophagus span a spectrum from asymptomatic motility disorders through mild achalasia to severe megaesophagus (83). Symptoms include dysphagia, odynophagia, esophageal reflux, weight loss, aspiration, cough, and regurgitation. As in idiopathic achalasia, the risk of esophageal carcinoma is elevated (13, 47). Megacolon is characterized by prolonged constipation and may give rise to fecaloma, volvulus, and bowel ischemia.

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

669

T. cruzi Infection in the Immunocompromised Host

DIAGNOSIS

Acute T. cruzi infection in organ transplantation recipients. Acute T. cruzi infection in organ recipients has several features that differ from those of acute T. cruzi infection in immunocompetent hosts. The incubation period can be prolonged: among the 15 patients for whom data were available in published reports, the mean time from transplantation to onset of symptoms of acute T. cruzi infection was 112 days (range, 23 to 420 days) (61, 66, 79, 99, 101, 157, 238, 279). A relatively severe clinical spectrum has been reported, with manifestations that included fever, malaise, anorexia, hepatosplenomegaly, acute myocarditis, and decreased cardiac function; two of the 18 reported patients presented with fulminant myocarditis and congestive heart failure (61, 279). Reactivation of chronic T. cruzi infection in organ recipients. Patients with chronic T. cruzi infection can be candidates for organ transplants. In a large cohort of heart transplant patients, survival of those who received the transplant because of chronic Chagas’ cardiomyopathy was longer than survival among those with idiopathic or ischemic cardiomyopathy, and T. cruzi reactivation was a rare cause of death (33, 38, 39). Reactivation should be considered in the differential diagnosis of febrile episodes and apparent rejection crises. In addition to fever and acute Chagas’ myocarditis in the transplanted heart, common manifestations of reactivation disease include inflammatory panniculitis and skin nodules (52, 102, 238). Central nervous system (CNS) involvement has been reported but is a much less frequent manifestation of reactivation among transplant recipients than in AIDS patients (5, 102, 180). Reactivation Chagas’ disease in HIV/AIDS patients. Reactivation of T. cruzi infection in HIV/AIDS patients can cause severe clinical disease with a high risk of mortality. However, as in organ transplant recipients, reactivation is not universal, even in those with low CD4⫹ lymphocyte counts. The only published prospective cohort study followed 53 HIV-T. cruzicoinfected patients in Brazil for 1 to 190 months; 11 (21%) had T. cruzi reactivation diagnosed based on symptoms and/or microscopically detectable parasitemia (260). Even among patients without clinical reactivation, the level of parasitemia is higher among HIV-coinfected than among HIV-negative patients (261). Symptomatic T. cruzi reactivation in AIDS patients is most commonly reported to cause meningoencephalitis and/or T. cruzi brain abscesses; the presentation may be confused with CNS toxoplasmosis and should be considered in the differential diagnosis of mass lesions on imaging or CNS syndromes in AIDS patients (70, 71, 88, 260). The second most commonly reported sign of reactivation is acute myocarditis, sometimes superimposed on preexisting chronic Chagas’ cardiomyopathy (260, 297). Patients may present with new arrhythmias, pericardial effusions, acute cardiac decompensation, or accelerated progression of existing chronic heart disease (100, 260). Acute meningoencephalitis and myocarditis can occur simultaneously. In the Brazilian cohort, cardiac reactivation was as frequent as CNS disease; cardiac manifestations of reactivated Chagas’ disease may pass undetected or mimic progression of chronic Chagas’ cardiomyopathy (260). Less common manifestations of reactivation in HIV/AIDS patients include skin lesions, erythema nodosum, and parasitic invasion of the peritoneum, stomach, or intestine (100, 261).

Appropriate diagnostic testing for T. cruzi infection varies depending on the phase of the disease and the status of the patient. In the United States, CDC provides consultation to health care providers concerning Chagas’ disease diagnostic testing (contact information is listed in “Antitrypanosomal Drugs” below).

Diagnosis of Acute T. cruzi Infection In the acute phase, motile trypomastigotes can be detected by microscopy of fresh preparations of anticoagulated blood or buffy coat (311). Parasites may also be visualized by microscopy of blood smears stained with Giemsa stain or other stains. Hemoculture in one of several types of standard parasitic medium (e.g., Novy-MacNeal-Nicolle) is relatively sensitive during the acute phase but requires 2 to 4 weeks to show replication. The level of parasitemia decreases within 90 days of infection, even without treatment, and becomes undetectable by microscopy in the chronic phase (306, 311). PCR is a sensitive diagnostic tool in the acute phase of Chagas’ disease and may also be used to monitor for acute T. cruzi infection in the recipient of an infected organ or after accidental exposure (133, 134, 157).

Diagnosis of Congenital T. cruzi Infection Early in life, congenital Chagas’ disease is an acute T. cruzi infection and similar diagnostic methods are employed. Concentration methods yield better sensitivity than direct examination of fresh blood. The microhematocrit method is the most widely used technique in Latin American health facilities. Fresh cord or neonatal blood is collected, sealed in four to six heparinized microhematocrit tubes, and centrifuged, and the buffy coat layer is examined by light microscopy (108). Parasitemia levels rise after birth and peak at or after 30 days of life (32). Repeated sampling on several occasions during the first months of life increases the sensitivity but may not be acceptable to parents (14, 32, 197). Hemoculture can increase sensitivity, but the technique is not widely available, and results are not available for 2 to 4 weeks. Molecular techniques have higher sensitivity and detect congenital infections earlier in life than the microhematocrit method (32, 92, 247). Transient detection of parasite DNA has occasionally been reported in specimens from infants who subsequently are found to be uninfected (32, 211). For this reason, a positive PCR on samples collected on two separate occasions may be used as a criterion for confirmation of congenital infection (32). PCR is increasingly used for the early diagnosis of congenital Chagas’ disease in Latin America and is the method of choice in industrialized countries (55, 140, 202, 247, 266). For infants not diagnosed at birth, conventional IgG serology (as outlined below for chronic T. cruzi infection) is recommended after 9 months of age, when transferred maternal antibody has disappeared and the congenital infection has passed into the chronic phase (32, 55, 56).

670

BERN ET AL.

CLIN. MICROBIOL. REV.

Diagnosis of Chronic T. cruzi infection

TREATMENT

Diagnosis of chronic infection relies on serological methods to detect IgG antibodies to T. cruzi, most commonly the enzyme-linked immunosorbent assay (ELISA) and immunofluorescent-antibody assay (IFA). No single assay has sufficient sensitivity and specificity to be relied on alone; two serological tests based on different antigens (e.g., whole parasite lysate and recombinant antigens) and/or techniques (e.g., ELISA, IFA, and immunoblotting) are used in parallel to increase the accuracy of the diagnosis (311). Inevitably, a proportion of individuals tested by two assays will have discordant serological results and need further testing to resolve their infection status. Specimens with positive results but low antibody titers are more likely to show discordance because results obtained by less sensitive tests may be negative. Published data suggest that the sensitivity of serological assays varies by geographical location, possibly due to T. cruzi strain differences and the resulting antibody responses (275, 293, 299). The status of some individuals remains difficult to resolve even after a third test, because there is no true gold standard assay for chronic T. cruzi infection (283). Assays such as the radioimmunoprecipitation assay (RIPA) and trypomastigote excreted-secreted antigen immunoblot (TESA-blot) are promoted as reference tests, but even these do not have perfect sensitivity and specificity and may not be capable of resolving the diagnosis (168, 272). Options for diagnostic T. cruzi serological testing are relatively limited in the United States. Several ELISA kits based on parasite lysate or recombinant antigens are Food and Drug Administration (FDA) cleared for diagnostic application. Use of an assay with validation data (e.g., a commercial kit shown to have acceptable sensitivity and specificity in a thorough study) is preferable to reliance on in-house tests for which no performance data are available (31).

Antitrypanosomal Drugs

Utility of PCR for Diagnosis or Monitoring PCR techniques provide the most sensitive tool to diagnose acute-phase and early congenital Chagas’ disease and to monitor for acute T. cruzi infection in the recipient of an infected organ or after accidental exposure (32, 65, 133). PCR assays usually show positive results days to weeks before circulating trypomastigotes are detectable on peripheral blood smears (267). Quantitative PCR assays (e.g., real-time PCR) are useful to monitor for reactivation in the immunosuppressed T. cruzi-infected host. In these patients, a positive result on conventional PCR does not prove reactivation, but quantitative PCR assays that indicate rising parasite numbers over time provide the earliest and most sensitive indicator of reactivation (89, 92). In chronic T. cruzi infection, PCR is used as a research tool but is not generally a useful diagnostic test. Although PCR results will be positive for a proportion of patients, the sensitivity is highly variable depending on the characteristics of the population tested, as well as the PCR primers and methods (25, 142, 314). For these reasons, negative results by PCR do not constitute evidence for lack of infection.

Nifurtimox and benznidazole are the only drugs with proven efficacy against Chagas’ disease (73, 177). Neither drug is approved by the U.S. FDA, but both can be obtained from the CDC and used under investigational protocols. Consultations and drug requests should be addressed to the Parasitic Diseases Public Inquiries line [(404) 718-4745; e-mail, [email protected]], the CDC Drug Service [(404) 639-3670], and, for emergencies after business hours and on weekends and federal holidays, the CDC Emergency Operations Center [(770) 488-7100]. Nifurtimox (Lampit, Bayer 2502), a nitrofuran, interferes with T. cruzi carbohydrate metabolism by inhibiting pyruvic acid synthesis. Gastrointestinal side effects are common, occurring in 30 to 70% of patients. These include anorexia leading to weight loss, nausea, vomiting, and abdominal discomfort. Neurological toxicity is also fairly common, including irritability, insomnia, disorientation, and, less often, tremors. Rare but more serious side effects include paresthesias, polyneuropathy, and peripheral neuritis. The peripheral neuropathy is dose dependent, appears late in the course of therapy, and should prompt interruption of treatment. Higher doses are often used in infants than in older children, and tolerance is better in children than in adults. Benznidazole (Rochagon, Roche 7-1051) is a nitroimidazole derivative, considered more trypanocidal than nifurtimox. Dermatological side effects are frequent, and consist of rashes due to photosensitization, rarely progressing to exfoliative dermatitis. Severe or exfoliative dermatitis or dermatitis associated with fever and lymphadenopathy should prompt immediate cessation of the drug. The peripheral neuropathy is dose dependent, usually occurs late in the course of therapy, and is an indication for immediate cessation of treatment; it is nearly always reversible but may take months to resolve. Bone marrow suppression is rare and should prompt immediate interruption of drug treatment. Patients should be monitored for dermatological side effects beginning 9 to 10 days after initiation of treatment. Benznidazole was well tolerated in two placebo-controlled trials with children (12% had a rash and ⬍5% had gastrointestinal symptoms in one study; ⬍10% had moderate reversible side effects in the other study) (7, 277). Side effects are more common in adults than in children. Treatment of Acute and Congenital T. cruzi infection In acute and early congenital Chagas’ disease, both drugs reduce the severity of symptoms, shorten the clinical course, and reduce the duration of detectable parasitemia (53, 306). The earliest trials of antitrypanosomal drugs were conducted with patients with acute Chagas’ disease in the 1960s and 1970s using nifurtimox (53, 306). Serological cure was documented at the 12-month follow-up in 81% of those treated in the acute phase (306). Treatment of Chronic T. cruzi Infection Until recently, only the acute phase, including early congenital infection, was thought to be responsive to antiparasitic

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

671

TABLE 6. Autochthonous human cases of Chagas’ disease in the United States Yr

State

Patient characteristics

1955

TX

1955 1982

TX CA

1983

TX

1998

TN

2006

LA

2006

TX

10-mo-old girl, acute Chagas’ disease, trypomastigotes on blood smear 2- to 3-wk-old boy, no details provided 56-yr-old woman with acute Chagas’ disease, trypomastigotes on blood smear 7-mo-old boy, fatal acute Chagas’ disease with myocarditis and pericardial effusion, postmortem diagnosis based on nests of T. cruzi in cardiac tissue 18-mo-old boy, febrile illness several wk after bug was found, positive T. cruzi PCR in multiple blood specimens 74-yr-old woman with history of triatomine bites but no symptoms of Chagas’ disease; positive IgG serology and T. cruzi hemoculture 12-mo-old boy with fever, large pericardial effusion and respiratory distress; trypomastigotes by microscopy in pericardial fluid

therapy. However, in the 1990s, 2 placebo-controlled trials of benznidazole treatment in children with chronic T. cruzi infection demonstrated approximately 60% cure as measured by conversion to negative serology 3 to 4 years after the end of treatment (7, 278). Several follow-up studies suggest that the earlier in life that children are treated, the higher the rate of reversion to negative serology (6, 281). Together with growing clinical experience across Latin America, these studies revolutionized management of children with Chagas’ disease, making early diagnosis and antitrypanosomal drug therapy the standard of care throughout the region (177, 311). There is currently a growing movement to offer treatment to older patients and those with early cardiomyopathy (30, 302, 311). In Latin America, most Chagas’ disease experts now believe that the majority of patients with chronic T. cruzi infection should be offered treatment, employing individual exclusion criteria such as an upper age limit of 50 or 55 years and the presence of advanced irreversible cardiomyopathy (276). This change in standards of practice is based in part on nonrandomized, nonblinded longitudinal studies that demonstrate decreased progression of Chagas’ cardiomyopathy and decreased mortality in adult patients treated with benznidazole (301, 302). A multicenter, randomized, placebo-controlled, double-blinded trial of benznidazole for patients with mild to moderate Chagas’ cardiomyopathy is under way and will help to clarify treatment efficacy for this group (http://clinicaltrials .gov/show/NCT00123916).

Evidence of autochthonous transmission

Reference(s)

Peridomestic infestation

342

No details provided Adult uninfected T. protracta found in house No vectors found, but household search was made in winter; house said to be in poor condition T. cruzi-infected T sanguisuga found in child’s crib

10 203, 265

Peridomestic and house T. sanguisuga infestation; 10/18 positive by T. cruzi PCR Mother uninfected; T. cruzi-infected T. gerstaeckeri collected near house

209 134 90 151; CDC, unpublished data

with standard courses of antitrypanosomal treatment; antiretroviral therapy should be optimized (143). EPIDEMIOLOGY OF CHAGAS’ DISEASE Since 1991, the estimated global prevalence of T. cruzi infection has fallen from 18 million to 8 million, due to intensive vector control and blood bank screening (87, 214). The Pan American Health Organization estimates that approximately 60,000 new T. cruzi infections occur each year (214). As other transmission routes have diminished, the proportion attributable to congenital infection has grown: an estimated 26% of incident infections now occur through mother-to-child transmission (214). In settings with endemic vector-borne transmission, T. cruzi infection is usually acquired in childhood. Because the infection is lifelong, the seroprevalence in an area with sustained vector-borne transmission rises with age, reflecting the cumulative incidence (98). Before widespread vector control was instituted in the early 1990s, it was common to find that ⬎60% of adults in an community where the disease was endemic were infected with T. cruzi (200, 230). In cross-sectional community surveys, most infected individuals are asymptomatic; an estimated 70 to 80% will remain asymptomatic throughout their lives (176, 234). Because cardiac and gastrointestinal manifestations usually begin in early adulthood and progress over a period of years to decades, the prevalence of clinical disease increases with increasing age (178).

Management of the Immunocompromised Host Antitrypanosomal treatment for reactivation in organ transplant recipients follows standard dosage regimens and promotes resolution of clinical symptoms and parasitemia. There are no data to indicate that prior treatment or posttransplant prophylaxis decreases the risk of reactivation; posttransplant prophylaxis is not routinely administered in heart transplant centers in Latin America (52). Antitrypanosomal therapy is thought to achieve a sterile cure in few, if any, adults with longstanding chronic infection, and treated patients should be considered to be at risk for reactivation. Reactivation in an HIV-coinfected patient should be treated

HUMAN CHAGAS’ DISEASE IN THE UNITED STATES Autochthonous Transmission to Humans Seven autochthonous vector-borne infections (four in Texas and one each in California, Tennessee, and Louisiana) have been reported since 1955 (Table 6) (10, 90, 134, 151, 209, 265, 342). Most reported cases have been in infants or small children; six of the seven infections were in the acute phase at the time of identification, and the diagnosis was sought because of symptoms and/or the presence of triatomine vectors. A survey conducted in the community of residence of the 1982 Califor-

672

BERN ET AL.

CLIN. MICROBIOL. REV. TABLE 7. Transfusion-related cases of Chagas’ disease in the United States Implicated blood component and donor origin

Yr

State

Recipient characteristics

1988

NY

1988

CA

1989

TX

1997

FL

2002

RI

11-yr-old girl with Hodgkin’s lymphoma, developed fever and pericarditis, trypomastigotes seen on blood smear; treated with nifurtimox and recovered 17-yr-old male post-bone marrow transplant with fulminant acute Chagas’ disease 59-yr-old female with metastatic colon cancer on chemotherapy, granulocytopenic, disseminated intravascular coagulation; developed fever, pulmonary infiltrates, bradycardia and atrioventricular block; parasites seen on bone marrow aspirate; died within 36 h of diagnosis 60-yr-old female with multiple myeloma; T. cruzi-infected donor unit detected during research study; recipient asymptomatic, treated with nifurtimox; died of underlying disease several yr later. 3-yr-old female with stage 4 neuroblastoma on chemotherapy, neutropenic, fever, trypomastigotes seen on blood smear; treated with nifurtimox but died of her underlying disease

nia case demonstrated positive complement fixation results in 6/241 (2.5%) residents tested (203). The rarity of autochthonous vector-borne transmission in the United States is assumed to result from better housing conditions that minimize vector-human contact. In addition, North American vectors may have lower transmission efficiency, due at least in part to delayed defecation (153, 203, 228). However, given that the vast majority of acute T. cruzi infections in immunocompetent individuals pass undiagnosed in Latin America, where the index of suspicion is much higher, undetected cases of autochthonous vector-borne transmission are presumed to occur.

Chagas’ Disease Burden among Latin American Immigrants The only direct assessments in Latin American populations living in the United States come from very limited local surveys and blood bank screening (see below) (42, 59, 148, 165, 167). No large representative surveys have ever been conducted, and blood bank data cannot be extrapolated with validity because donors are not representative of the larger population. The only recent data come from a survey of Latin American immigrants attending churches in Los Angeles County; a total of 10 (1%) of 985 adults tested had positive results by serological testing (290). Based on the reported number of immigrants from countries in Latin America where Chagas’ disease is endemic and the estimated national T. cruzi seroprevalences in their countries of origin, there are an estimated 300,000 persons with T. cruzi infection currently living in the United States (29). Patients with clinical manifestations of Chagas’ disease, especially cardiomyopathy, are assumed to be present but largely unrecognized in hospitals and health care facilities in the United States, but systematic data are sparse (126). Recent targeted studies in a Los Angeles hospital demonstrated positive results by T. cruzi serological tests among 15 (16%) of 93 Latin American patients with a diagnosis of idiopathic cardiomyopathy and 11 (4.6%) of 239 patients with conduction system abnormalities on ECG and at least 1 year of residence in Latin America (190, 291).

Reference

Platelets, Bolivia

114

Not specified, Mexico

113

Unknown; had received ⬎500 units, including red blood cells and platelets

67

Platelets, Bolivia

166

Platelets, Bolivia

351

Blood-Borne Transmission and Blood Donor Screening A total of 5 transfusion-associated T. cruzi infections have been documented in the United States since the late 1980s (Table 7) (59, 67, 114, 166, 351). All infected recipients had underlying malignancies and were immunosuppressed. Platelet units from Bolivian donors were implicated in 3 of 5 cases. Several patients had severe manifestations of Chagas’ disease, including acute myocarditis, acute atrioventricular block, severe congestive heart failure, pericarditis with T. cruzi in the pericardial fluid, and possible meningoencephalitis (67, 114, 351). The recipient of a platelet unit detected as infected during a research study had T. cruzi infection detected by PCR and serology during prospective monitoring but never developed symptoms (166). In December 2006, the FDA approved an ELISA to screen for antibodies to T. cruzi in donated blood (59). The radioimmunoprecipitation assay (RIPA) has been used as the confirmatory test (1, 31, 257). The American Red Cross and Blood Systems Inc. voluntarily began screening all blood donations in January 2007, and in subsequent months, many other blood centers starting screening as well. As of 2 September 2011, 1,459 confirmed seropositive donations have been detected in 43 states, with the largest numbers found in California, Florida, and Texas (1). A second T. cruzi antibody screening test was approved in April 2010. In December 2010, FDA issued specific guidance for appropriate use of the screening tests (103). Current FDA recommendations are to screen all blood donors initially, and if a donor’s sample tests negative using one of the two FDA-approved screening tests, no testing of future donations by that donor is necessary. No supplemental test has been approved, and donors are deferred indefinitely on the basis of positive screening test results alone. This strategy will be reviewed by FDA at upcoming meetings of the Blood Products Advisory Committee; the risk of newly acquired blood donor infections, including results from longitudinal studies of repeat blood donors, will be considered. Screening of blood donations remains voluntary, although most blood centers are currently following FDA recommendations. In data from the first 16 months of screening, comprising

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

673

TABLE 8. Published reports of organ transplant-derived cases of Chagas’ disease in the United Statesa Yr

State of organ harvest

2001

GA

2001

Implicated organ

Recipient characteristics and outcome

Reference

El Salvador

Kidney-pancreas

61

GA

El Salvador

Kidney

2001

GA

El Salvador

Liver

2005

CA

US-born (mother from Mexico)

Heart

2006

CA

El Salvador

Heart

37-yr-old female with fever 6 wk posttransplant and T. cruzi on blood smear, died of Chagas’ myocarditis 7 mo posttransplant despite prolonged course of nifurtimox 69-yr-old female, asymptomatic, T. cruzi hemoculture positive; diagnosis sought because of recipient 1 above; treated with nifurtimox, survived 32-yr-old female, asymptomatic, T. cruzi hemoculture positive; diagnosis sought because of recipient 1 above; treated with nifurtimox but died of unrelated causes 64-yr-old male with anorexia, fever, diarrhea; diagnosed with organ rejection, treated with steroids; 8 wk posttransplant T. cruzi found on blood smear; PCRs became negative on nifurtimox; died of rejection 20 wk posttransplant 73-yr-old male with fever, fatigue, rash, T. cruzi on blood smear 7 wk posttransplant; parasitemia cleared with nifurtimox; switched to benznidazole because of tremors; died of heart failure 25 wk posttransplant

a

Donor origin

61 61 157

157

Three additional unpublished cases are known to have occurred (2 heart transplants and 1 liver transplant).

⬎14 million blood donations, the overall seroprevalence was 1:27,500 based on donations screened, with the highest rates in Florida (1:3,800), followed by California (1:8,300) (31). Because large blood donor studies prior to FDA approval of the screening ELISA were conducted in southern California with permanent deferral of all repeatedly reactive donors, a substantial number of infected individuals were already removed from the local donor pool, and the reported prevalence in California is thought to represent an underestimate (42, 59, 165, 167). From preliminary data, 29 (28%) of 104 T. cruziinfected donors were born in Mexico, 27 (26%) in the United States, 17 (16%) in El Salvador, and 11 (11%) in Bolivia; the remaining 20 donors were born in 9 other countries of Central and South America (31). Among confirmed infected donors born in the United States, 10 individuals reported no specific risk factors for T. cruzi infection. All of these donors reported outdoor activities (e.g., hunting, camping, or extensive gardening) in the southern United States, which may indicate potential autochthonous exposure to the vector or animal reservoirs.

curred. All three of these recipients were treated and survived their T. cruzi infection (65; S. Huprikar and B. Kubak, unpublished data). When an infected organ donor is detected, recipient monitoring relies primarily on detection of the parasite by microscopy, culture, and/or PCR, because seroconversion may be delayed or never occur in immunocompromised individuals (65, 238). Molecular techniques usually show positive results days to weeks before circulating trypomastigotes are visible by microscopy of peripheral blood. Transplant-transmitted T. cruzi infection may have a longer incubation period than vector-borne infection; parasitemia is usually detected within 2 to 3 months, but the delay can be as long as 6 months. A frequently recommended monitoring schedule consists of weekly specimens for 2 months, specimens every 2 weeks up to 4 months, and then monthly specimens afterwards (65, 238). In the absence of other indications and assuming no evidence of infection has been detected, the monitoring interval can be lengthened after 6 months posttransplantation.

Organ Donor-Derived Transmission and Organ Donor Screening

Unanswered Questions and Priorities for Research and Programs

A total of five instances of organ-derived transmission from three donors are documented in the published literature in the United States (Table 8) (60, 61, 157). Four of the five recipients died. One patient died from acute Chagas’ myocarditis; T. cruzi infection was not the primary cause of but may have contributed to the other deaths (61, 157). In all of these instances of transmission, donor infections were not suspected until at least one recipient presented with symptomatic acute Chagas’ disease (60, 61, 157). More recently, some organ procurement organizations have begun selective or universal screening of donated organs (65). Three transmission events (in two heart recipients in 2006 and 2010 and one liver recipient in 2006) were detected through systematic laboratory monitoring when their respective donors were identified as infected shortly after the transplants oc-

The United States faces important public health challenges for the prevention, control, and management of T. cruzi infection and Chagas’ disease (86). Patients with undiagnosed Chagas’ cardiomyopathy go unrecognized, impeding their optimal management. The large number of undetected T. cruzi infections sustains the risk of transmission through blood and organ donation and from mother to child. Currently, obstetricians have limited knowledge of congenital T. cruzi transmission risk, and almost no screening of at-risk women is carried out (48). Many health care providers in all specialties fail to consider the diagnosis of Chagas’ disease in patients at risk and are unaware that antitrypanosomal treatment is available (280, 300); the possibility that treatment could decrease the risk of progression of disease in infected individuals is therefore not realized. Worldwide, programs to control Chagas’ disease are ham-

674

BERN ET AL.

CLIN. MICROBIOL. REV.

pered by the lack of adequate tools, and these challenges are equally salient in the United States (283). Point-of-care diagnostic tests would allow physicians to make a rapid diagnosis in patients in whom Chagas’ disease is suspected and provide a practical means to identify women at risk of transmitting the infection to their infants. However, the sensitivity of current T. cruzi rapid tests shows wide geographic variation (275, 299); there is a need for screening tests with high sensitivity, especially for T. cruzi infections originating in geographic areas such as Mexico and the United States, where current tests appear to have low sensitivity (275; CDC, unpublished data). Two other diagnostic needs are critical: a practical, timely test of cure and indicators to distinguish patients who are likely to develop clinical disease from those likely to remain asymptomatic. Unfortunately, neither of these tools is currently on the horizon. Pediatric formulations of existing drugs are of immediate concern and expected to be available soon (91). However, new treatment drugs with high efficacy and better safety profiles, especially in adults, are needed (295). To inform effective policy for Chagas’ disease control in the United States, significant gaps in our knowledge must also be addressed. Systematic, rigorous population-based data to determine infection prevalence and morbidity are needed to inform prevention strategies. Pilot studies in hospitals with a high proportion of women born in Latin America would help to define practical methods to target screening for congenital transmission. More thorough identification of the T. cruzi strains circulating in the United States will add to our assessment of transfusion risk and understanding of the molecular epidemiology of the disease (164). More comprehensive assessment of the magnitude of local transmission risk and the factors influencing vector and reservoir host distribution and human contact are important to inform control efforts. Improved knowledge of the local epidemiology and ecology will allow more efficient, effective targeting of limited resources and raise awareness of Chagas’ disease in the United States. As improved control of vector- and blood-borne T. cruzi transmission decreases the burden in countries where the disease is historically endemic and imported Chagas’ disease is increasingly recognized outside Latin America, the United States— which confronts the challenges faced both by countries where the disease is endemic and by those where it is not—can play an important role in addressing the altered epidemiology of Chagas’ disease in the 21st century. REFERENCES 1. AABB Chagas Biovigilance Network. 2011, posting date. Reports through 04/18/2011. http://www.aabb.org/programs/biovigilance/Pages/chagas.aspx. 2. Abad-Franch, F., and F. A. Monteiro. 2007. Biogeography and evolution of Amazonian triatomines (Heteroptera: Reduviidae): implications for Chagas disease surveillance in humid forest ecoregions. Mem. Inst. Oswaldo Cruz 102(Suppl. 1):57–70. 3. Acquatella, H. 2007. Echocardiography in Chagas heart disease. Circulation 115:1124–1131. 4. Aguilar, H. M., F. Abad-Franch, J. C. Dias, A. C. Junqueira, and J. R. Coura. 2007. Chagas disease in the Amazon region. Mem. Inst. Oswaldo Cruz 102(Suppl. 1):47–56. 5. Altclas, J., et al. 2005. Chagas disease in bone marrow transplantation: an approach to preemptive therapy. Bone Marrow Transplant. 36:123–129. 6. Andrade, A. L., et al. 2004. Benznidazole efficacy among Trypanosoma cruzi-infected adolescents after a six-year follow-up. Am. J. Trop. Med. Hyg. 71:594–597. 7. Andrade, A. L., et al. 1996. Randomised trial of efficacy of benznidazole in treatment of early Trypanosoma cruzi infection. Lancet 348:1407–1413. 8. Andrade, M. C., et al. 2009. Nonspecific lymphocytic myocarditis in ba-

9. 10. 11.

12. 13.

14.

15.

16.

17.

18. 19.

20.

21.

22. 23. 24.

25.

26. 27.

28.

29. 30. 31.

32. 33.

34. 35.

36.

37.

38.

boons is associated with Trypanosoma cruzi infection. Am. J. Trop. Med. Hyg. 81:235–239. Andrade, S. G. 1982. The influence of the strain of Trypanosoma cruzi in placental infections in mice. Trans. R. Soc. Trop. Med. Hyg. 76:123–128. Anonymous. 1956. Found: two cases of Chagas disease. Texas Health Bull. 9:11–13. Arganaraz, E. R., et al. 2001. Blood-sucking lice may disseminate Trypanosoma cruzi infection in baboons. Rev. Inst. Med. Trop. Sao Paulo 43:271– 276. Arnold, H. L., and D. B. Bell. 1944. Kissing bug bites. J. Allergy Clin. Immunol. 74:436–442. Atias, A. 1994. A case of congenital chagasic megaesophagus: evolution until death caused by esophageal neoplasm, at 27 years of age. Rev. Med. Chil. 122:319–322. Azogue, E., and C. Darras. 1991. Prospective study of Chagas disease in newborn children with placental infection caused by Trypanosoma cruzi (Santa Cruz-Bolivia). Rev. Soc. Bras. Med. Trop. 24:105–109. Badra, E. S., et al. 2008. Histopathological changes in the placentas and fetuses of mice infected with Trypanosoma cruzi isolated from the Myotis nigricans nigricans bat. J. Comp. Pathol. 139:108–112. Barcan, L., et al. 2005. Transmission of T. cruzi infection via liver transplantation to a nonreactive recipient for Chagas’ disease. Liver Transplant. 11:1112–1116. Barnabe, C., R. Yaeger, O. Pung, and M. Tibayrenc. 2001. Trypanosoma cruzi: a considerable phylogenetic divergence indicates that the agent of Chagas disease is indigenous to the native fauna of the United States. Exp. Parasitol. 99:73–79. Barr, S., D. Baker, and J. Markovits. 1986. Trypanosomiasis and laryngeal paralysis in a dog. J. Am. Vet. Med. Assoc. 188:1307–1309. Barr, S. C., C. C. Brown, V. A. Dennis, and T. R. Klei. 1991. The lesions and prevalence of Trypanosoma cruzi in opossums and armadillos from southern Louisiana. J. Parasitol. 77:624–627. Barr, S. C., V. A. Dennis, and T. R. Klei. 1991. Serologic and blood culture survey of Trypanosoma cruzi infection in four canine populations of southern Louisiana. Am. J. Vet. Res. 52:570–573. Barr, S. C., K. A. Gossett, and T. R. Klei. 1991. Clinical, clinicopathologic, and parasitologic observations of trypanosomiasis in dogs infected with North American Trypanosoma cruzi isolates. Am. J. Vet. Res. 52:954–960. Barr, S. C., et al. 1989. Chronic dilatative myocarditis caused by Trypanosoma cruzi in two dogs. J. Am. Vet. Med. Assoc. 195:1237–1241. Barr, S. C., et al. 1995. Trypanosoma cruzi infection in Walker hounds from Virginia. Am. J. Vet. Res. 56:1037–1044. Basombrio, M. A., et al. 1999. The transmission de Chagas disease in Salta and the detection of congenital cases. Medicina (Buenos Aires) 59(Suppl. 2):143–146. Basquiera, A. L., et al. 2003. Risk progression to chronic Chagas cardiomyopathy: influence of male sex and of parasitaemia detected by PCR. Heart 89:1186–1190. Beard, C. B., et al. 2003. Chagas disease in a domestic transmission cycle, southern Texas, USA. Emerg. Infect. Dis. 9:103–105. Beard, C. B., D. G. Young, J. F. Butler, and D. A. Evans. 1988. First isolation of Trypanosoma cruzi from a wild-caught Triatoma sanguisuga (LeConte) (Hemiptera: Triatominae) in Florida, USA. J. Parasitol. 74:343– 344. Beltrao, B., et al. 2009. Investigation of two outbreaks of suspected oral transmission of acute Chagas disease in the Amazon region, Para State, Brazil, in 2007. Trop. Doct. 39:231–232. Bern, C., and S. P. Montgomery. 2009. An estimate of the burden of Chagas disease in the United States. Clin. Infect. Dis. 49:e52–54. Bern, C., et al. 2007. Evaluation and treatment of Chagas disease in the United States: a systematic review. JAMA 298:2171–2181. Bern, C., S. P. Montgomery, L. Katz, S. Caglioti, and S. L. Stramer. 2008. Chagas disease and the US blood supply. Curr. Opin. Infect. Dis. 21:476– 482. Bern, C., et al. 2009. Congenital Trypanosoma cruzi transmission in Santa Cruz, Bolivia. Clin. Infect. Dis. 49:1667–1674. Bestetti, R. B., and T. A. Theodoropoulos. 2009. A systematic review of studies on heart transplantation for patients with end-stage Chagas’ heart disease. J. Card. Fail. 15:249–255. Bittencourt, A. L. 1992. Possible risk factors for vertical transmission of Chagas’ disease. Rev. Inst. Med. Trop. Sao Paulo 34:403–408. Bittencourt, A. L., L. A. Rodrigues de Freitas, M. O. Galvao de Araujo, and K. Jacomo. 1981. Pneumonitis in congenital Chagas’ disease. A study of ten cases. Am. J. Trop. Med. Hyg. 30:38–42. Bittencourt, A. L., M. Sadigursky, and H. S. Barbosa. 1975. Congenital Chagas’ disease. Study of 29 cases. Rev. Inst. Med. Trop. Sao Paulo 17: 146–159. Bittencourt, A. L., G. O. Vieira, H. C. Tavares, E. Mota, and J. Maguire. 1984. Esophageal involvement in congenital Chagas’ disease. Report of a case with megaesophagus. Am. J. Trop. Med. Hyg. 33:30–33. Bocchi, E. A., and A. Fiorelli. 2001. The Brazilian experience with heart

VOL. 24, 2011

39.

40. 41.

42.

43.

44.

45.

46.

47.

48.

49.

50. 51.

52. 53.

54.

55.

56.

57. 58.

59.

60.

61.

62.

63. 64.

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

transplantation: a multicenter report. J. Heart Lung Transplant. 20:637– 645. Bocchi, E. A., and A. Fiorelli. 2001. The paradox of survival results after heart transplantation for cardiomyopathy caused by Trypanosoma cruzi. First Guidelines Group for Heart Transplantation of the Brazilian Society of Cardiology. Ann. Thorac. Surg. 71:1833–1838. Bosseno, M. F., et al. 2002. Predominance of Trypanosoma cruzi lineage I in Mexico. J. Clin. Microbiol. 40:627–632. Bradley, K. K., D. K. Bergman, J. P. Woods, J. M. Crutcher, and L. V. Kirchhoff. 2000. Prevalence of American trypanosomiasis (Chagas disease) among dogs in Oklahoma. J. Am. Vet. Med. Assoc. 217:1853–1857. Brashear, R. J., et al. 1995. Detection of antibodies to Trypanosoma cruzi among blood donors in the southwestern and western United States. I. Evaluation of the sensitivity and specificity of an enzyme immunoassay for detecting antibodies to T. cruzi. Transfusion 35:213–218. Briones, M. R., R. P. Souto, B. S. Stolf, and B. Zingales. 1999. The evolution of two Trypanosoma cruzi subgroups inferred from rRNA genes can be correlated with the interchange of American mammalian faunas in the Cenozoic and has implications to pathogenicity and host specificity. Mol. Biochem. Parasitol. 104:219–232. Brisse, S., C. Barnabe, and M. Tibayrenc. 2000. Identification of six Trypanosoma cruzi phylogenetic lineages by random amplified polymorphic DNA and multilocus enzyme electrophoresis. Int. J. Parasitol. 30:35–44. Brisse, S., et al. 2003. Evidence for genetic exchange and hybridization in Trypanosoma cruzi based on nucleotide sequences and molecular karyotype. Infect. Genet. Evol. 2:173–183. Brown, E. L., et al. 2010. Seroprevalence of Trypanosoma cruzi among eleven potential reservoir species from six states across the southern United States. Vector Borne Zoonotic Dis. 10:757–763. Brucher, B. L., H. J. Stein, H. Bartels, H. Feussner, and J. R. Siewert. 2001. Achalasia and esophageal cancer: incidence, prevalence, and prognosis. World J. Surg. 25:745–749. Buekens, P., et al. 2008. Mother-to-child transmission of Chagas’ disease in North America: why don’t we do more? Matern. Child Health J. 12:283– 286. Burkholder, J. E., T. C. Allison, and V. P. Kelly. 1980. Trypanosoma cruzi (Chagas) (Protozoa: Kinetoplastida) in invertebrate, reservoir, and human hosts of the lower Rio Grande valley of Texas. J. Parasitol. 66:305–311. Burleigh, B. A., and N. W. Andrews. 1995. The mechanisms of Trypanosoma cruzi invasion of mammalian cells. Annu. Rev. Microbiol. 49:175–200. Campbell, D. A., S. J. Westenberger, and N. R. Sturm. 2004. The determinants of Chagas disease: connecting parasite and host genetics. Curr. Mol. Med. 4:549–562. Campos, S. V., et al. 2008. Risk factors for Chagas’ disease reactivation after heart transplantation. J. Heart Lung Transplant. 27:597–602. Cancado, J. R., and Z. Brener. 1979. Terapeutica, p. 362–424. In Z. Brener and Z. Andrade (ed.), Trypanosoma cruzi e doenc¸a de Chagas, 1st ed. Guanabara Koogan, Rio de Janeiro, Brazil. Cantey, P. T., S. Hand, M. Currier, P. Jett, and S. Montgomery. 2008. Chagas disease in Mississippi: investigation of suspected autochthonous infections in the United States, abstr. H3, p. 146. 2008 Int. Conf. Emerg. Infect. Dis. International Conference on Emerging Infectious Diseases, Atlanta, GA. Carlier, Y., and F. Torrico. 2003. Congenital infection with Trypanosoma cruzi: from mechanisms of transmission to strategies for diagnosis and control. Rev. Soc. Bras. Med. Trop. 36:767–771. Carlier, Y., and C. Truyens. 2010. Maternal-fetal transmission of Trypanosoma cruzi, p. 539–581. In J. Telleria and M. Tibayrenc (ed.), American trypanosomiasis-Chagas disease: one hundred years of research. Elsevier, New York, NY. Carranza, J. C., et al. 2009. Trypanosoma cruzi maxicircle heterogeneity in Chagas disease patients from Brazil. Int. J. Parasitol. 39:963–973. Castanera, M. B., M. A. Lauricella, R. Chuit, and R. E. Gurtler. 1998. Evaluation of dogs as sentinels of the transmission of Trypanosoma cruzi in a rural area of north-western Argentina. Ann. Trop. Med. Parasitol. 92: 671–683. Centers for Disease Control and Prevention. 2007. Blood donor screening for Chagas disease—United States, 2006–2007. MMWR Morb. Mortal. Wkly. Rep. 56:141–143. Centers for Disease Control and Prevention. 2006. Chagas disease after organ transplantation—Los Angeles, California, 2006. MMWR Morb. Mortal. Wkly. Rep. 55:798–800. Centers for Disease Control and Prevention. 2002. Chagas disease after organ transplantation—United States, 2001. MMWR Morb. Mortal. Wkly. Rep. 51:210–212. Chagas, C. 1909. Nova tripanosomiaze humana. Estudos sobre morfolojia e o ciclo evolutivo do Schizotrypanum cruzi n.g., n. sp., ajente etiolojico de nova entidade morbida do homen. Mem. Inst. Oswaldo Cruz 1:159–218. Chagas, C. 1924. Sobre a verificac¸˜ao do “Trypanosoma cruzi” em macacos do Para´ (Chrysothrix sciureus). Sci. Me´d. 2:75–76. Chagas, C. 1912. Sobre um trypanosomo do tatu ´, Tatusia novemcincta, transmittido pela Triatoma geniculata Latr. (1811). Possibilidade de ser o

65.

66.

67. 68.

69. 70.

71. 72.

73. 74.

75.

76.

77.

78.

79. 80. 81.

82.

83.

84.

85. 86. 87.

88.

89.

90. 91. 92.

93. 94.

675

tatu ´ um depositario do Trypanosoma cruzi no mundo exterior Brasil. Me´dico I:305–306. Chin-Hong, P. V., et al. 2011. Screening and treatment of Chagas disease in organ transplant recipients in the United States: recommendations from the Chagas in Transplant Working Group. Am. J. Transplant. 11:672–680. Chocair, P. R., E. Sabbaga, V. Amato Neto, M. Shiroma, and G. M. de Goes. 1981. Kidney transplantation: a new way of transmitting Chagas disease. Rev. Inst. Med. Trop. Sao Paulo 23:280–282. Cimo, P. L., W. E. Luper, and M. A. Scouros. 1993. Transfusion-associated Chagas’ disease in Texas: report of a case. Tex. Med. 89:48–50. Clark, C. G., and O. J. Pung. 1994. Host specificity of ribosomal DNA variation in sylvatic Trypanosoma cruzi from North America. Mol. Biochem. Parasitol. 66:175–179. Cohen, J. E., and R. E. Gurtler. 2001. Modeling household transmission of American trypanosomiasis. Science 293:694–698. Cordova, E., A. Boschi, J. Ambrosioni, C. Cudos, and M. Corti. 2008. Reactivation of Chagas disease with central nervous system involvement in HIV-infected patients in Argentina, 1992–2007. Int. J. Infect. Dis. 12:587– 592. Cordova, E., E. Maiolo, M. Corti, and T. Orduna. 2010. Neurological manifestations of Chagas’ disease. Neurol. Res. 32:238–244. Corrales, R. M., et al. 2009. Congenital Chagas disease involves Trypanosoma cruzi sub-lineage IId in the northwestern province of Salta, Argentina. Infect. Genet. Evol. 9:278–282. Coura, J. R., and S. L. de Castro. 2002. A critical review on Chagas disease chemotherapy. Mem. Inst. Oswaldo Cruz 97:3–24. Coura, J. R., A. C. Junqueira, O. Fernandes, S. A. Valente, and M. A. Miles. 2002. Emerging Chagas disease in Amazonian Brazil. Trends Parasitol. 18:171–176. Covarrubias, C., M. Cortez, D. Ferreira, and N. Yoshida. 2007. Interaction with host factors exacerbates Trypanosoma cruzi cell invasion capacity upon oral infection. Int. J. Parasitol. 37:1609–1616. Cura, C. I., et al. 2010. Trypanosoma cruzi I genotypes in different geographical regions and transmission cycles based on a microsatellite motif of the intergenic spacer of spliced-leader genes. Int. J. Parasitol. 40:1599– 1607. Davis, D. J. 1943. Triatoma sanguisuga (LeConte) and Triatoma ambigua Neiva as natural carriers of Trypanosoma cruzi in Texas. Public Health Rep. 58:353–354. Davis, D. S., L. H. Russell, L. G. Adams, R. G. Yaeger, and R. M. Robinson. 1980. An experimental infection of Trypanosoma cruzi in striped skunks (Mephitis mephitis). J. Wildl. Dis. 16:403–406. de Faria, J. B., and G. Alves. 1993. Transmission of Chagas’ disease through cadaveric renal transplantation. Transplantation 56:1583–1584. de Freitas, J. M., et al. 2006. Ancestral genomes, sex, and the population structure of Trypanosoma cruzi. PLoS Pathog. 2:e24. Delgado, M. A., and C. A. Santos-Buch. 1978. Transplacental transmission and fetal parasitosis of Trypanosoma cruzi in outbred white Swiss mice. Am. J. Trop. Med. Hyg. 27:1108–1115. de Noya, B. A., et al. 2010. Large urban outbreak of orally acquired acute Chagas disease at a school in Caracas, Venezuela. J. Infect. Dis. 201:1308– 1315. de Oliveira, R. B., L. E. Troncon, R. O. Dantas, and U. G. Menghelli. 1998. Gastrointestinal manifestations of Chagas’ disease. Am. J. Gastroenterol. 93:884–889. de Rezende, J. M., and H. Moreira. 1988. Chagasic megaesophagus and megacolon. Historical review and present concepts. Arq. Gastroenterol. 25(Spec No.):32–43. DeShazo, T. 1943. A survey of Trypanosoma cruzi infection in Triatoma spp. collected in Texas. J. Bacteriol. 46:219–220. Dias, J. C. 2009. Elimination of Chagas disease transmission: perspectives. Mem. Inst. Oswaldo Cruz 104(Suppl. 1):41–45. Dias, J. C., A. C. Silveira, and C. J. Schofield. 2002. The impact of Chagas disease control in Latin America: a review. Mem. Inst. Oswaldo Cruz 97:603–612. Diazgranados, C. A., et al. 2009. Chagasic encephalitis in HIV patients: common presentation of an evolving epidemiological and clinical association. Lancet Infect. Dis. 9:324–330. Diez, M., et al. 2007. Usefulness of PCR strategies for early diagnosis of Chagas’ disease reactivation and treatment follow-up in heart transplantation. Am. J. Transplant. 7:1633–1640. Dorn, P. L., et al. 2007. Autochthonous transmission of Trypanosoma cruzi, Louisiana. Emerg. Infect. Dis. 13:605–607. Drugs for Neglected Disease Initiative. 2010, posting date. Chagas disease: DNDi strategy. http://www.treatchagas.org/rd_dndi_strategy.aspx. Duffy, T., et al. 2009. Accurate real-time PCR strategy for monitoring bloodstream parasitic loads in Chagas disease patients. PLoS Negl. Trop. Dis. 3:e419. Eads, R. B., and B. G. Hightower. 1952. Blood parasites of south Texas rodents. J. Parasitol. 38:89–90. Eads, R. B., H. A. Trevino, and E. G. Campos. 1963. Triatoma (Hemiptera:

676

95. 96.

97. 98. 99.

100.

101. 102. 103.

104. 105.

106. 107. 108.

109.

110.

111. 112.

113.

114. 115.

116.

117.

118. 119. 120.

121. 122.

123.

124.

BERN ET AL. Reduviidae) infected with Trypanosoma cruzi in south Texas wood rat dens. Southwest Nat. 8:38–42. Ekkens, D. 1981. Nocturnal flights of Triatoma (Hemiptera: Reduviidae) in Sabino Canyon, Arizona. I. Light collections. J. Med. Entomol. 18:211–227. Ekkens, D. 1984. Nocturnal flights of Triatoma (Hemiptera: Reduviidae) in Sabino Canyon, Arizona. II. Neotoma lodge studies. J. Med. Entomol 21:140–144. Estrada-Franco, J. G., et al. 2006. Human Trypanosoma cruzi infection and seropositivity in dogs, Mexico. Emerg. Infect. Dis. 12:624–630. Feliciangeli, M. D., et al. 2003. Chagas disease control in Venezuela: lessons for the Andean region and beyond. Trends Parasitol. 19:44–49. Ferraz, A. S., and J. F. Figueiredo. 1993. Transmission of Chagas’ disease through transplanted kidney: occurrence of the acute form of the disease in two recipients from the same donor. Rev. Inst. Med. Trop. Sao Paulo 35:461–463. Ferreira, M. S., et al. 1997. Reactivation of Chagas’ disease in patients with AIDS: report of three new cases and review of the literature. Clin. Infect. Dis. 25:1397–1400. Figueiredo, J. F., et al. 1990. Transmission of Chagas disease through renal transplantation: report of a case. Trans. R. Soc. Trop. Med. Hyg. 84:61–62. Fiorelli, A. I., et al. 2005. Later evolution after cardiac transplantation in Chagas’ disease. Transplant. Proc. 37:2793–2798. Food and Drug Administration. December 2010, posting date. Guidance for industry: use of serological tests to reduce the risk of transmission of Trypanosoma cruzi infection in whole blood and blood components intended for transfusion. http://www.fda.gov/BiologicsBloodVaccines /GuidanceComplianceRegulatoryInformation/Guidances/default.htm. Fores, R., et al. 2007. Chagas disease in a recipient of cord blood transplantation. Bone Marrow Transplant. 39:127–128. Fox, J. C., S. A. Ewing, R. G. Buckner, D. Whitenack, and J. H. Manley. 1986. Trypanosoma cruzi infection in a dog from Oklahoma. J. Am. Vet. Med. Assoc. 189:1583–1584. Frazier, C. A. 1974. Biting insect survey: a statistical report. Ann. Allergy 32:200–204. Freilij, H., and J. Altcheh. 1995. Congenital Chagas’ disease: diagnostic and clinical aspects. Clin. Infect. Dis. 21:551–555. Freilij, H., L. Muller, and S. M. Gonzalez Cappa. 1983. Direct micromethod for diagnosis of acute and congenital Chagas’ disease. J. Clin. Microbiol. 18:327–330. Freitas, J. L. P., et al. 1952. Primeiras verficac¸o ˜es de transmissa˜o acidental da mole´stia de Chagas ao homem por transfusa˜o de sangue. Rev. Paul. Med. 40:36–40. Galvao, C., R. Carcavallo, S. Rocha Dda, and J. Jurberg. 2003. A checklist of the current valid species of the subfamily Triatominae Jeannel, 1919 (Hemiptera, Reduviidae, Triatominae) and their geographical distribution with nomenclatural and taxonomic notes. Zootaxa 202:1–36. Gascon, J., C. Bern, and M. J. Pinazo. 2010. Chagas disease in Spain, the United States and other non-endemic countries. Acta Trop. 115:22–27. Gaunt, M., and M. Miles. 2000. The ecotopes and evolution of triatomine bugs (triatominae) and their associated trypanosomes. Mem. Inst. Oswaldo Cruz 95:557–565. Gerseler, P. J., J. I. Ito, B. R. Tegtmeier, P. R. Kerndt, and R. Krance. 1988. Fulminant Chagas disease in bone marrow transplantation, abstr. 418. Abstr. 27th Intersci. Conf. Antimicrob. Agents Chemother., Washington, DC. Grant, I. H., et al. 1989. Transfusion-associated acute Chagas disease acquired in the United States. Ann. Intern. Med. 111:849–851. Grieves, J. L., et al. 2008. Trypanosoma cruzi in non-human primates with a history of stillbirths: a retrospective study (Papio hamadryas spp.) and case report (Macaca fascicularis). J. Med. Primatol. 37:318–328. Griffith, M. E. 1948. The bloodsucking conenose, or “big bedbug,” Triatoma sanguisuga (Leconte), in an Oklahoma City household. Proc. Oklahoma Acad. Sci. 28:24–27. Grijalva, M. J., et al. 2003. Seroprevalence and risk factors for Trypanosoma cruzi infection in the Amazon region of Ecuador. Am. J. Trop. Med. Hyg. 69:380–385. Groce, B. 2008. Trypanosoma cruzi in wild raccoons and opossums from Kentucky. Western Kentucky University, Bowling Green, KY. Grogl, M., R. E. Kuhn, D. S. Davis, and G. E. Green. 1984. Antibodies to Trypanosoma cruzi in coyotes in Texas. J. Parasitol. 70:189–191. Grundemann, A. W. 1947. Studies on the biology of the Triatoma sanguisuga (Leconte) in Kansas (Reduviidae: Triatominae). Kansas Entomol. Soc. 20:77–85. Guedes, P. M., et al. 2002. The dog as model for chemotherapy of the Chagas’ disease. Acta Trop. 84:9–17. Gurtler, R. E., et al. 2007. Domestic dogs and cats as sources of Trypanosoma cruzi infection in rural northwestern Argentina. Parasitology 134: 69–82. Gurtler, R. E., et al. 1991. Chagas disease in north-west Argentina: infected dogs as a risk factor for the domestic transmission of Trypanosoma cruzi. Trans. R. Soc. Trop. Med. Hyg. 85:741–745. Gurtler, R. E., et al. 1998. Influence of humans and domestic animals on the

CLIN. MICROBIOL. REV.

125.

126. 127.

128.

129.

130. 131. 132.

133. 134.

135.

136.

137.

138.

139.

140. 141. 142.

143.

144. 145.

146.

147. 148.

149.

150. 151.

household prevalence of Trypanosoma cruzi in Triatoma infestans populations in northwest Argentina. Am. J. Trop. Med. Hyg. 58:748–758. Haberkorn, A., and R. Gonnert. 1972. Animal experimental investigation into the activity of nifurtimox against Trypanosoma cruzi. Arzneimittelforschung 22:1570–1582. Hagar, J. M., and S. H. Rahimtoola. 1991. Chagas’ heart disease in the United States. N. Engl. J. Med. 325:763–768. Hall, C. A., E. M. Pierce, A. N. Wimsatt, T. Hobby-Dolbeer, and J. B. Meers. 2010. Virulence and vertical transmission of two genotypically and geographically diverse isolates of Trypanosoma cruzi in mice. J. Parasitol. 96: 371–376. Hall, C. A., C. Polizzi, M. J. Yabsley, and T. M. Norton. 2007. Trypanosoma cruzi prevalence and epidemiologic trends in lemurs on St. Catherines Island, Georgia. J. Parasitol. 93:93–96. Hancock, K., et al. 2005. Prevalence of antibodies to Trypanosoma cruzi in raccoons (Procyon lotor) from an urban area of northern Virginia. J. Parasitol. 91:470–472. Herman, C. M., and J. I. Bruce. 1962. Occurrence of Trypanosoma cruzi in Maryland. Proc. Helminthol. Soc. Washington 29:55–58. Herrer, A. 1964. Chagas’ disease in Peru. I. The epidemiological importance of the guinea pig. Trop. Geogr. Med. 16:146–151. Herrera, C., et al. 2007. Identifying four Trypanosoma cruzi I isolate haplotypes from different geographic regions in Colombia. Infect. Genet. Evol. 7:535–539. Herwaldt, B. L. 2001. Laboratory-acquired parasitic infections from accidental exposures. Clin. Microbiol. Rev. 14:659–688. Herwaldt, B. L., et al. 2000. Use of PCR to diagnose the fifth reported US case of autochthonous transmission of Trypanosoma cruzi, in Tennessee, 1998. J. Infect. Dis. 181:395–399. Hwang, W. S., G. Zhang, D. Maslov, and C. Weirauch. 2010. Infection rates of Triatoma protracta (Uhler) with Trypanosoma cruzi in Southern California and molecular identification of trypanosomes. Am. J. Trop. Med. Hyg. 83:1020–1022. Hypsa, V., et al. 2002. Phylogeny and biogeography of Triatominae (Hemiptera: Reduviidae): molecular evidence of a New World origin of the Asiatic clade. Mol. Phylogenet. Evol. 23:447–457. Ibarra-Cerdena, C. N., V. Sanchez-Cordero, A. Townsend Peterson, and J. M. Ramsey. 2009. Ecology of North American triatominae. Acta Trop. 110:178–186. Ikenga, J. O., and J. V. Richerson. 1984. Trypanosoma cruzi (Chagas) (Protozoa: Kinetoplastida: Trypanosomatidae) in invertebrate and vertebrate hosts from Brewster County in Trans-Pecos Texas. J. Econ. Entomol. 77:126–129. Iwagami, M., et al. 2007. Molecular phylogeny of Trypanosoma cruzi from Central America (Guatemala) and a comparison with South American strains. Parasitol. Res. 102:129–134. Jackson, Y., et al. 2009. Congenital transmission of Chagas disease in Latin American immigrants in Switzerland. Emerg. Infect. Dis. 15:601–603. John, D. T., and K. L. Hoppe. 1986. Trypanosoma cruzi from wild raccoons in Oklahoma. Am. J. Vet. Res. 47:1056–1059. Junqueira, A. C., E. Chiari, and P. Wincker. 1996. Comparison of the PCR with two classical parasitological methods for the diagnosis of Chagas disease in an endemic region of north-eastern Brazil. Trans. R. Soc. Trop. Med. Hyg. 90:129–132. Kaplan, J. E., et al. 2009. Guidelines for prevention and treatment of opportunistic infections in HIV-infected adults and adolescents: recommendations from CDC, the National Institutes of Health, and the HIV Medicine Association of the Infectious Diseases Society of America. MMWR Recommend. Rep. 58:1–207. Karsten, V., C. Davis, and R. Kuhn. 1992. Trypanosoma cruzi in wild raccoons and opossums in North Carolina. J. Parasitol. 78:547–549. Kasa, T. J., G. D. Lathrop, H. J. Dupuy, C. H. Bonney, and J. D. Toft. 1977. An endemic focus of Trypanosoma cruzi infection in a subhuman primate research colony. J. Am. Vet. Med. Assoc. 171:850–854. Kierszenbaum, F., C. A. Gottlieb, and D. B. Budzko. 1981. Antibodyindependent, natural resistance of birds to Trypanosoma cruzi infection. J. Parasitol. 67:656–660. Kimball, B. M. 1894. Conorhinus sanguisuga, its habits and life history. Kansas Acad. Sci. 14:128–131. Kirchhoff, L. V., A. A. Gam, and F. C. Gilliam. 1987. American trypanosomiasis (Chagas’ disease) in Central American immigrants. Am. J. Med. 82:915–920. Kjos, S. A. 2009. Characterization of Chagas disease transmission in peridomestic settings in the southwestern United States, p. 55. 59th Annu. James H. Steele Conf. Dis. Nat. Transm. Man, Ft. Worth, TX. Diseases in Nature Conference, Austin, TX. http://www.dshs.state.tx.us/idcu/health /zoonosis/education/conference/DIN/2009/. Kjos, S. A., et al. 2008. Distribution and characterization of canine Chagas disease in Texas. Vet. Parasitol. 152:249–256. Kjos, S. A., K. F. Snowden, and J. K. Olson. 2009. Biogeography and Trypanosoma cruzi infection prevalence of Chagas disease vectors in Texas, USA. Vector Borne Zoonotic Dis. 9:41–50.

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

152. Klotz, J. H., et al. 2010. “Kissing bugs”: potential disease vectors and cause of anaphylaxis. Clin. Infect. Dis. 50:1629–1634. 153. Klotz, S. A., et al. 2009. Feeding behavior of triatomines from the southwestern United States: an update on potential risk for transmission of Chagas disease. Acta Trop. 111:114–118. 154. Kofoid, C. A., and F. Donat. 1933. The experimental transfer of Trypanosoma cruzi from naturally infected Triatoma protracta to mammals in California. Univ. Calif. Publ. Zool. 86:257–259. 155. Kofoid, C. A., and L. McCulloch. 1916. On Trypanosoma triatomae, a new flagellate from a hemipteran bug from the nests of the wood rat Neotoma fuscipes. Univ. Calif. Publ. Zool. 16:113–126. 156. Kofoid, C. A., and B. G. Whitaker. 1936. Natural infection of American human trypanosomiasis in two species of cone-nosed bugs, Triatoma protracta Uhler and Triatoma uhleri Neiva, in the western United States. J. Parasitol. 22:259–263. 157. Kun, H., et al. 2009. Transmission of Trypanosoma cruzi by heart transplantation. Clin. Infect. Dis. 48:1534–1540. 158. Lambert, R. C., K. N. Kolivras, L. M. Resler, C. C. Brewster, and S. L. Paulson. 2008. The potential for emergence of Chagas disease in the United States. Geospatial Health 2:227–239. 159. Lane, R. S., et al. 1999. Anti-arthropod saliva antibodies among residents of a community at high risk for Lyme disease in California. Am. J. Trop. Med. Hyg. 61:850–859. 160. Lathrop, G. D., and A. J. Ominsky. 1965. Chagas disease study in a group of individuals bitten by North American triatomids. Aeromed. Rev. 9:1–5. 161. Leconte, J. L. 1855. Remarks on two species of American Cimex. Proc. Acad. Nat. Sci. Philadelphia 7:404. 162. Legey, A. P., A. P. Pinho, S. C. Chagas Xavier, L. L. Leon, and A. M. Jansen. 1999. Humoral immune response kinetics in Philander opossum and Didelphis marsupialis infected and immunized by Trypanosoma cruzi employing an immunofluorescence antibody test. Mem. Inst. Oswaldo Cruz 94:371–376. 163. Legey, A. P., et al. 2003. Trypanosoma cruzi in marsupial didelphids (Philander frenata and Didelphis marsupialis): differences in the humoral immune response in natural and experimental infections. Rev. Soc. Bras. Med. Trop. 36:241–248. 164. Leiby, D. A., R. M. Herron, Jr., G. Garratty, and B. L. Herwaldt. 2008. Trypanosoma cruzi parasitemia in US blood donors with serologic evidence of infection. J. Infect. Dis. 198:609–613. 165. Leiby, D. A., R. M. Herron, Jr., E. J. Read, B. A. Lenes, and R. J. Stumpf. 2002. Trypanosoma cruzi in Los Angeles and Miami blood donors: impact of evolving donor demographics on seroprevalence and implications for transfusion transmission. Transfusion 42:549–555. 166. Leiby, D. A., B. A. Lenes, M. A. Tibbals, and M. T. Tames-Olmedo. 1999. Prospective evaluation of a patient with Trypanosoma cruzi infection transmitted by transfusion. N. Engl. J. Med. 341:1237–1239. 167. Leiby, D. A., et al. 1997. Seroepidemiology of Trypanosoma cruzi, etiologic agent of Chagas’ disease, in US blood donors. J. Infect. Dis. 176:1047–1052. 168. Leiby, D. A., et al. 2000. Serologic testing for Trypanosoma cruzi: comparison of radioimmunoprecipitation assay with commercially available indirect immunofluorescence assay, indirect hemagglutination assay, and enzyme-linked immunosorbent assay kits. J. Clin. Microbiol. 38:639–642. 169. Lent, H., and P. Wygodzinsky. 1979. Revision of the Triatominae (Hemiptera, Reduviidae), and their significance as vectors of Chagas’ disease. Bull. Am. Museum Nat. History 63:123–520. 170. Levy, M. Z., et al. 2006. Peri-urban infestation by Triatoma infestans carrying Trypanosoma cruzi in Arequipa, Peru. Emerg. Infect. Dis. 12:1345–1352. 171. Lewis, M. D., et al. 2009. Genotyping of Trypanosoma cruzi: systematic selection of assays allowing rapid and accurate discrimination of all known lineages. Am. J. Trop. Med. Hyg. 81:1041–1049. 172. Llewellyn, M. S., et al. 2009. Trypanosoma cruzi IIc: phylogenetic and phylogeographic insights from sequence and microsatellite analysis and potential impact on emergent Chagas disease. PLoS Negl. Trop. Dis. 3:e510. 173. Llewellyn, M. S., et al. 2009. Genome-scale multilocus microsatellite typing of Trypanosoma cruzi discrete typing unit I reveals phylogeographic structure and specific genotypes linked to human infection. PLoS Pathog. 5:e1000410. 174. Macedo, A. M., and S. D. J. Pena. 1998. Genetic variability of Trypanosoma cruzi: implications for the pathogenesis of Chagas disease. Parasitol. Today 14:119–124. 175. Machado, C. A., and F. J. Ayala. 2001. Nucleotide sequences provide evidence of genetic exchange among distantly related lineages of Trypanosoma cruzi. Proc. Natl. Acad. Sci. U. S. A. 98:7396–7401. 176. Magill, A. J., and S. G. Reed. 2000. American trypanosomiasis, p. 653–664. In G. T. Strickland (ed.), Hunter’s tropical medicine and emerging diseases, 8th ed. W.B. Saunders Company, Philadelphia, PA. 177. Maguire, J. H. 2004. Trypanosoma, p. 2327–2334. In S. Gorbach, J. Bartlett, and N. Blacklow (ed.), Infectious diseases, 2nd ed. Lippincott, Williams & Wilkins, Philadelphia, PA. 178. Maguire, J. H., et al. 1987. Cardiac morbidity and mortality due to Chagas’

179. 180.

181.

182. 183.

184.

185.

186. 187.

188.

189.

190.

191.

192.

193.

194. 195. 196.

197.

198.

199.

200.

201.

202.

203. 204. 205.

206.

677

disease: prospective electrocardiographic study of a Brazilian community. Circulation 75:1140–1145. Maloney, J., et al. 2010. Seroprevalence of Trypanosoma cruzi in raccoons from Tennessee. J. Parasitol. 96:353–358. Marchiori, P. E., et al. 2007. Late reactivation of Chagas’ disease presenting in a recipient as an expansive mass lesion in the brain after heart transplantation of chagasic myocardiopathy. J. Heart Lung Transplant. 26:1091– 1096. Marcili, A., et al. 2009. Comparative phylogeography of Trypanosoma cruzi TcIIc: new hosts, association with terrestrial ecotopes, and spatial clustering. Infect. Genet. Evol. 9:1265–1274. Marin-Neto, J. A., E. Cunha-Neto, B. C. Maciel, and M. V. Simoes. 2007. Pathogenesis of chronic Chagas heart disease. Circulation 115:1109–1123. Martinez-Ibarra, J. A., et al. 2007. Biology of three species of North American Triatominae (Hemiptera: Reduviidae: Triatominae) fed on rabbits. Mem. Inst. Oswaldo Cruz 102:925–930. Martinez-Ibarra, J. A., L. Galaviz-Silva, C. L. Campos, and J. C. TrujilloGarcia. 1992. Distribucion de los triatominos asociados al domicilio humano en el municipio de general Teran, Nuevo Leon, Mexico. Southwest Entomol. 17:261–264. McKeever, S., G. W. Gorman, and L. Norman. 1958. Occurrence of a Trypanosoma cruzi-like organism in some mammals from southwestern Georgia and northwestern Florida. J. Parasitol. 44:583–587. McKown, R. D., S. J. Upton, R. D. Klemm, and R. K. Ridley. 1990. New host and locality record for Trypanosoma peromysci. J. Parasitol. 76:281–283. Mehringer, P. J., and S. F. Wood. 1958. A resampling of wood rat houses and human habitations in Griffith Park, Los Angeles, for Triatoma protracta and Trypanosoma cruzi. Bull. South. Calif. Acad. Sci. 57:39–46. Meirelles, M. N., and W. De Souza. 1985. Killing of Trypanosoma cruzi and Leishmania mexicana, and survival of Toxoplasma gondii, in chicken macrophages in vitro. J. Submicrosc. Cytol. 17:327–334. Meurs, K. M., M. A. Anthony, M. Slater, and M. W. Miller. 1998. Chronic Trypanosoma cruzi infection in dogs: 11 cases (1987–1996). J. Am. Vet. Med. Assoc. 213:497–500. Meymandi, S. K., et al. 2009. Prevalence of Chagas disease in U.S. immigrant population with conduction abnormalities on electrocardiogram, abstr. 410. Abstr. 58th Annu. Meet. Am. Soc. Trop. Med. Hyg., Washington, DC, 18 to 22 November 2009. Miles, M. A., et al. 1981. Do radically dissimilar Trypanosoma cruzi strains (zymodemes) cause Venezuelan and Brazilian forms of Chagas’ disease? Lancet i:1338–1340. Miles, M. A., M. D. Feliciangeli, and A. R. de Arias. 2003. American trypanosomiasis (Chagas’ disease) and the role of molecular epidemiology in guiding control strategies. BMJ 326:1444–1448. Miles, M. A., et al. 2009. The molecular epidemiology and phylogeography of Trypanosoma cruzi and parallel research on Leishmania: looking back and to the future. Parasitology 136:1509–1528. Ministe´rio da Sau ´de Brasil. 2005. Brazilian consensus on Chagas disease. Rev. Soc. Bras. Med. Trop. 38(Suppl. 3):7–29. Moffett, A., et al. 2009. A global public database of disease vector and reservoir distributions. PLoS Negl. Trop. Dis. 3:e378. Moncayo, A., and M. I. Ortiz Yanine. 2006. An update on Chagas disease (human American trypanosomiasis). Ann. Trop. Med. Parasitol. 100:663– 677. Mora, M. C., et al. 2005. Early diagnosis of congenital Trypanosoma cruzi infection using PCR, hemoculture, and capillary concentration, as compared with delayed serology. J. Parasitol. 91:1468–1473. Mortenson, E. W., and J. D. Walsh. 1963. Review of the Triatoma protracta problem in the Sierra Nevada foothills of California, p. 44–45. In Proceedings of the 31st Annual Conference of the California Mosquito Control Association. California Mosquito Control Association, Sacramento, CA. Mota, E., et al. 1984. Megaesophagus and seroreactivity to Trypanosoma cruzi in a rural community in northeast Brazil. Am. J. Trop. Med. Hyg. 33:820–826. Mott, K. E., J. S. Lehman, Jr., R. hoff, et al. 1976. The epidemiology and household distribution of seroreactivity to Trypanosoma cruzi in a rural community in northeast Brazil. Am. J. Trop. Med. Hyg. 25:552–562. Mott, K. E., et al. 1978. House construction, triatomine distribution, and household distribution of seroreactivity to Trypanosoma cruzi in a rural community in northeast Brazil. Am. J. Trop. Med. Hyg. 27:1116–1122. Mun ˜ oz, J., M. Portus, M. Corachan, V. Fumado, and J. Gascon. 2007. Congenital Trypanosoma cruzi infection in a non-endemic area. Trans. R. Soc. Trop. Med. Hyg. 101:1161–1162. Navin, T. R., et al. 1985. Human and sylvatic Trypanosoma cruzi infection in California. Am. J. Public Health 75:366–369. Nichols, N., and T. W. Green. 1963. Allergic reactions to “kissing bug” bites. Calif. Med. 98:267–268. Nieto, P. D., et al. 2009. Comparison of two immunochromatographic assays and the indirect immunofluorescence antibody test for diagnosis of Trypanosoma cruzi infection in dogs in south central Louisiana. Vet. Parasitol. 165:241–247. Nissen, E. E., E. L. Roberson, L. B. Liham, and W. L. Hanson. 1977.

678

207. 208. 209.

210.

211.

212.

213.

214.

215.

216. 217. 218. 219.

220.

221.

222.

223.

224. 225.

226.

227. 228.

229.

230.

231.

232.

BERN ET AL. Naturally occurring Chagas disease in a South Carolina puppy. 114th Am. Assoc. Vet. Parasitol. Forum, p. 122. Noble, E. R., and D. Shipman. 1958. Trypanosomes in American ground squirrels. J. Eukaryot. Microbiol. 5:247–249. Nobrega, A. A., et al. 2009. Oral transmission of Chagas disease by consumption of acai palm fruit, Brazil. Emerg. Infect. Dis. 15:653–655. Ochs, D. E., V. S. Hnilica, D. R. Moser, J. H. Smith, and L. V. Kirchhoff. 1996. Postmortem diagnosis of autochthonous acute chagasic myocarditis by PCR amplification of a species-specific DNA sequence of Trypanosoma cruzi. Am. J. Trop. Med. Hyg. 54:526–529. O’Connor, O., M. F. Bosseno, C. Barnabe, E. J. Douzery, and S. F. Breniere. 2007. Genetic clustering of Trypanosoma cruzi I lineage evidenced by intergenic miniexon gene sequencing. Infect. Genet. Evol. 7:587–593. Oliveira, I., F. Torrico, J. Munoz, and J. Gascon. 2010. Congenital transmission of Chagas disease: a clinical approach. Expert Rev. Anti Infect. Ther. 8:945–956. Olsen, P. F., J. P. Shoemaker, H. F. Turner, and K. L. Hays. 1964. Incidence of Trypanosoma cruzi (Chagas) in wild vectors and reservoirs in east-central Alabama. J. Parasitol. 50:599–603. Olson, L. C., S. F. Skinner, J. L. Palotay, and G. E. McGhee. 1986. Encephalitis associated with Trypanosoma cruzi in a Celebes black macaque. Lab. Anim. Sci. 36:667–670. Organizacio ´n Panamericana de la Salud. 2006. Estimacio ´n cuantitativa de la enfermedad de Chagas en las Americas OPS/HDM/CD/425-06. Organizacio ´n Panamericana de la Salud, Washington, DC. Packchanian, A. 1940. Experimental transmission of Trypanosoma cruzi infection in animals by Triatoma sanguisuga ambigua. Public Health Rep. 55:1526–1532. Packchanian, A. 1943. The infectivity of the Texas strain of Trypansoma cruzi to man. Am. J. Trop. Med. 23:309–314. Packchanian, A. 1939. Natural infection of Triatoma gerstaeckeri with Trypanosoma cruzi in Texas. Public Health Rep. 54:1547–1554. Packchanian, A. 1940. Natural infection of Triatoma heidemanni with Trypanosoma cruzi in Texas. Public Health Rep. 55:1300–1306. Packchanian, A. 1942. Reservoir hosts of Chagas’ disease in the State of Texas. Natural infection of nine-banded armadillo (Dasypus novemcinctus texanus), house mice (Mus musculus), opossum (Didelphis virginiana), and wood rats (Neotoma micropus micropus), with Trypanosoma cruzi in the state of Texas. Am. J. Trop. Med. 22:623–631. Pan American Health Organization. 2010, posting date. Chagas disease (American trypanosomiasis). http://www.paho.org/english/ad/dpc/cd/chagas .htm. Paredes, G. E. A., J. Valdez Miranda, B. Nogueda Torres, R. AlejandreAguilar, and R. Canett Romero. 2001. Vectorial importance of Triatominae bugs (Hemiptera: Reduviidae) in Guaymas, Mexico. Rev. Latinoam. Microbiol. 43:119–122. Parrish, E. A., and A. J. Mead. 2010. Determining the prevalence of Trypanosoma cruzi in road-killed opossums (Didelphis virginiana) from Baldwin County, Georgia, using PCR. Georgia J. Sci. 68:132–139. Patterson, J. S., C. J. Schofield, J. P. Dujardin, and M. A. Miles. 2001. Population morphometric analysis of the tropicopolitan bug Triatoma rubrofasciata and relationships with old world species of Triatoma: evidence of New World ancestry. Med. Vet. Entomol. 15:443–451. Pietrzak, S. M., and O. J. Pung. 1998. Trypanosomiasis in raccoons from Georgia. J. Wildl. Dis. 34:132–136. Pinho, A. P., E. Cupolillo, R. H. Mangia, O. Fernandes, and A. M. Jansen. 2000. Trypanosoma cruzi in the sylvatic environment: distinct transmission cycles involving two sympatric marsupials. Trans. R. Soc. Trop. Med. Hyg. 94:509–514. Pinnas, J. L., R. E. Lindberg, T. M. Chen, and G. C. Meinke. 1986. Studies of kissing bug-sensitive patients: evidence for the lack of cross-reactivity between Triatoma protracta and Triatoma rubida salivary gland extracts. J. Allergy Clin. Immunol. 77:364–370. Pinto, C. M., et al. 2010. Using museum collections to detect pathogens. Emerg. Infect. Dis. 16:356–357. Pippin, W. F. 1970. The biology and vector capability of Triatoma sanguisuga texana Usinger and Triatoma gerstaeckeri (Stal) compared with Rhodnius prolixus (Stal) (Hemiptera: Triatominae). J. Med. Entomol. 7:30–45. Pippin, W. F., P. F. Law, and M. J. Gaylor. 1968. Triatoma sanguisuga texana Usinger and Triatoma sanguisuga indictiva Neiva naturally infected with Trypanosoma cruzi Chagas in Texas (Hemiptera: Triatominae) (Kinetoplastida: Trypanosomidae). J. Med. Entomol. 5:134. Pless, M., et al. 1992. The epidemiology of Chagas’ disease in a hyperendemic area of Cochabamba, Bolivia: a clinical study including electrocardiography, seroreactivity to Trypanosoma cruzi, xenodiagnosis, and domiciliary triatomine distribution. Am. J. Trop. Med. Hyg. 47:539–546. Pung, O. J., C. W. Banks, D. N. Jones, and M. W. Krissinger. 1995. Trypanosoma cruzi in wild raccoons, opossums, and triatomine bugs in southeast Georgia, USA. J. Parasitol. 81:324–326. Pung, O. J., J. Spratt, C. G. Clark, T. M. Norton, and J. Carter. 1998. Trypanosoma cruzi infection of free-ranging lion-tailed macaques (Macaca

CLIN. MICROBIOL. REV.

233. 234. 235.

236. 237. 238. 239. 240.

241.

242.

243.

244.

245.

246. 247.

248. 249.

250.

251. 252. 253.

254.

255.

256.

257.

258.

259. 260.

261.

262.

silenus) and ring-tailed lemurs (Lemur catta) on St. Catherine’s Island, Georgia, USA. J. Zoo Wildl. Med. 29:25–30. Rassi, A., Jr., A. Rassi, and W. C. Little. 2000. Chagas’ heart disease. Clin. Cardiol. 23:883–889. Rassi, A., Jr., A. Rassi, and J. A. Marin-Neto. 2010. Chagas disease. Lancet 375:1388–1402. Rassi, A., Jr., A. Rassi, and S. G. Rassi. 2007. Predictors of mortality in chronic Chagas disease: a systematic review of observational studies. Circulation 115:1101–1108. Rassi, A., Jr., S. G. Rassi, and A. Rassi. 2001. Sudden death in Chagas’ disease. Arq. Bras. Cardiol. 76:75–96. Reisenman, C. E., et al. 2010. Infection of kissing bugs with Trypanosoma cruzi, Tucson, Arizona, USA. Emerg. Infect. Dis. 16:400–405. Riarte, A., et al. 1999. Chagas’ disease in patients with kidney transplants: 7 years of experience 1989–1996. Clin. Infect. Dis. 29:561–567. Roellig, D. M., et al. 2008. Molecular typing of Trypanosoma cruzi isolates, United States. Emerg. Infect. Dis. 14:1123–1125. Roellig, D. M., A. E. Ellis, and M. J. Yabsley. 2009. Genetically different isolates of Trypanosoma cruzi elicit different infection dynamics in raccoons (Procyon lotor) and Virginia opossums (Didelphis virginiana). Int. J. Parasitol. 39:1603–1610. Roellig, D. M., A. E. Ellis, and M. J. Yabsley. 2009. Oral transmission of Trypanosoma cruzi with opposing evidence for the theory of carnivory. J. Parasitol. 95:360–364. Roellig, D. M., et al. 2010. Experimental infection of two South American reservoirs with four distinct strains of Trypanosoma cruzi. Parasitology 137: 959–966. Rohr, A. S., N. A. Marshall, and A. Saxon. 1984. Successful immunotherapy for Triatoma protracta-induced anaphylaxis. J. Allergy Clin. Immunol. 73: 369–375. Romana, C. A., D. Brunstein, A. Collin-Delavaud, O. Sousa, and E. OrtegaBarria. 2003. Public policies of development in Latin America and Chagas’ disease. Lancet 362:579. Rosypal, A. C., R. R. Tidwell, and D. S. Lindsay. 2007. Prevalence of antibodies to Leishmania infantum and Trypanosoma cruzi in wild canids from South Carolina. J. Parasitol. 93:955–957. Rowland, M. E., et al. 2010. Factors associated with Trypanosoma cruzi exposure among domestic canines in Tennessee. J. Parasitol. 96:547–551. Russomando, G., et al. 1998. Treatment of congenital Chagas’ disease diagnosed and followed up by the PCR. Am. J. Trop. Med. Hyg. 59:487– 491. Ryan, C. P., P. E. Hughes, and E. B. Howard. 1985. American trypanosomiasis (Chagas’ disease) in a striped skunk. J. Wildl. Dis. 21:175–176. Ryckman, R. E. 1962. Biosystematics and hosts of the Triatoma protracta complex in North America (Hemiptera: Reduviidae) (Rodentia: Cricetidae). Univ. Calif. Publ. Entomol. 27:93–240. Ryckman, R. E. 1965. Epizootiology of Trypanosoma cruzi in southwestern North America. I. New collection records and hosts for Trypanosoma cruzi Chagas (Kinetoplastida: Trypanosomidae) (Hemiptera: Triatominae). J. Med. Entomol. 2:87–89. Ryckman, R. E. 1971. The genus Paratriatoma in western North America. J. Med. Entomol. 8:87–97. Ryckman, R. E. 1981. The kissing bug problem in western North America. Bull. Soc. Vector Ecol. 6:167–169. Ryckman, R. E. 1954. Lizards: a laboratory host for Triatominae and Trypansoma cruzi Chagas (Hemiptera: Reduviidae) (Protomonadida: Trypanosomidae). Trans. Am. Microsc. Soc. 63:215–218. Ryckman, R. E. 1984. The Triatominae of North and Central America and the West Indies: a checklist with synonymy (Hemiptera: Reduviidae: Triatominae). Bull. Soc. Vector Ecol. 9:71–83. Ryckman, R. E. 1986. The vertebrate hosts of the Triatominae of North and Central America and the West Indies (Hemiptera: Reduviidae: Triatominae). Bull. Soc. Vector Ecol. 11:221–241. Ryckman, R. E., and J. V. Ryckman. 1967. Epizootiology of Trypanosoma cruzi in southwestern North America. XII. Does Gause’s rule apply to the ectoparasitic Triatominae? (Hemiptera: Reduviidae) (Kinetoplastidae: Trypanosomidae) (Rodentia: Cricetidae). J. Med. Entomol 4:379–386. Sabino, E. C., et al. 2010. Enhanced classification of Chagas serologic results and epidemiologic characteristics of seropositive donors at three large blood centers in Brazil. Transfusion 50:2628–2637. Samudio, F., E. Ortega-Barria, A. Saldana, and J. Calzada. 2007. Predominance of Trypanosoma cruzi I among Panamanian sylvatic isolates. Acta Trop. 101:178–181. Sarkar, S., et al. 2010. Chagas disease risk in Texas. PLoS Negl. Trop. Dis. 4:e836. Sartori, A. M., et al. 2007. Manifestations of Chagas disease (American trypanosomiasis) in patients with HIV/AIDS. Ann. Trop. Med. Parasitol. 101:31–50. Sartori, A. M., et al. 2002. Trypanosoma cruzi parasitemia in chronic Chagas disease: comparison between human immunodeficiency virus (HIV)-positive and HIV-negative patients. J. Infect. Dis. 186:872–875. Schaffer, G. D., W. L. Hanson, W. R. Davidson, and V. F. Nettles. 1978.

VOL. 24, 2011

263.

264.

265.

266.

267.

268.

269. 270. 271.

272.

273.

274.

275.

276.

277.

278.

279.

280. 281.

282.

283.

284. 285.

286.

287.

288.

289.

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

Hematotropic parasites of translocated raccoons in the southeast. J. Am. Vet. Med. Assoc. 173:1148–1151. Schenone, H., M. Gaggero, J. Sapunar, M. C. Contreras, and A. Rojas. 2001. Congenital Chagas disease of second generation in Santiago, Chile. Report of two cases. Rev. Inst. Med. Trop. Sao Paulo 43:231–232. Schielke, J. E., R. Selvarangan, K. B. Kyes, and T. R. Fritsche. 2002. Laboratory diagnosis of Trypanosoma cruzi infection in a colony-raised pigtailed macaque. Contemp. Top. Lab. Anim. Sci. 41:42–45. Schiffler, R. J., G. P. Mansur, T. R. Navin, and K. Limpakarnjanarat. 1984. Indigenous Chagas’ disease (American trypanosomiasis) in California. JAMA 251:2983–2984. Schijman, A. G. 2006. Congenital Chagas disease, p. 223–259. In I. K. Mushahwar (ed.), Congenital and other related infectious diseases of the newborn, vol. 13. Elsevier, Amsterdam, Netherlands. Schijman, A. G., et al. 2000. Early diagnosis of recurrence of Trypanosoma cruzi infection by PCR after heart transplantation of a chronic Chagas’ heart disease patient. J. Heart Lung Transplant. 19:1114–1117. Schmunis, G. A. 1991. Trypanosoma cruzi, the etiologic agent of Chagas’ disease: status in the blood supply in endemic and nonendemic countries. Transfusion 31:547–557. Schmunis, G. A., and J. R. Cruz. 2005. Safety of the blood supply in Latin America. Clin. Microbiol. Rev. 18:12–29. Schofield, C. J., and C. Galvao. 2009. Classification, evolution, and species groups within the Triatominae. Acta Trop. 110:88–100. Secretaria de Vigilancia em Saude de Brasil. 2007, posting date. Doenca de Chagas Aguda. Nota Tecnica, 9 de Outubro de 2007. http://portal.saude .gov.br/portal/arquivos/pdf/nota_chagas_091007.pdf. Silveira-Lacerda, E. P., et al. 2004. Chagas’ disease: application of TESAblot in inconclusive sera from a Brazilian blood bank. Vox Sang. 87:204– 207. Sjogren, R. D., and R. E. Ryckman. 1966. Epizootiology of Trypanosoma cruzi in southwestern North America. 8. Nocturnal flights of Triatoma protracta (Uhler) as indicated by collections at black light traps (Hemiptera: Reduviidae: Triatominae). J. Med. Entomol. 3:81–92. Snider, T. G., R. G. Yaeger, and J. Dellucky. 1980. Myocarditis caused by Trypanosoma cruzi in a native Louisiana dog. J. Am. Vet. Med. Assoc. 177:247–249. Sosa-Estani, S., et al. 2008. Use of a rapid test on umbilical cord blood to screen for Trypanosoma cruzi infection in pregnant women in Argentina, Bolivia, Honduras, and Mexico. Am. J. Trop. Med. Hyg. 79:755–759. Sosa-Estani, S., and E. L. Segura. 2006. Etiological treatment in patients infected by Trypanosoma cruzi: experiences in Argentina. Curr. Opin. Infect. Dis. 19:583–587. Sosa-Estani, S., and E. L. Segura. 1999. Treatment of Trypanosoma cruzi infection in the undetermined phase. Experience and current guidelines of treatment in Argentina. Mem. Inst. Oswaldo Cruz 94(Suppl. 1):363–365. Sosa Estani, S., et al. 1998. Efficacy of chemotherapy with benznidazole in children in the indeterminate phase of Chagas’ disease. Am. J. Trop. Med. Hyg. 59:526–529. Souza, F. F., et al. 2008. Acute chagasic myocardiopathy after orthotopic liver transplantation with donor and recipient serologically negative for Trypanosoma cruzi: a case report. Transplant. Proc. 40:875–878. Stimpert, K. K., and S. P. Montgomery. 2010. Physician awareness of Chagas disease, USA. Emerg. Infect. Dis. 16:871–872. Streiger, M. L., M. L. del Barco, D. L. Fabbro, E. D. Arias, and N. A. Amicone. 2004. Longitudinal study and specific chemotherapy in children with chronic Chagas’ disease, residing in a low endemicity area of Argentina. Rev. Soc. Bras. Med. Trop. 37:365–375. Sullivan, T. D., T. McGregor, R. B. Eads, and D. J. Davis. 1949. Incidence of Trypanosoma cruzi, Chagas, in Triatoma (Hemiptera: Reduviidae) in Texas. Am. J. Trop. Med. 29:453–458. Tarleton, R. L., R. Reithinger, J. A. Urbina, U. Kitron, and R. E. Gurtler. 2007. The challenges of Chagas disease—grim outlook or glimmer of hope. PLoS Med. 4:e332. Telford, S. R., Jr., and D. J. Forrester. 1991. Hemoparasites of raccoons (Procyon lotor) in Florida. J. Wildl. Dis. 27:486–490. Theis, J. H., M. Tibayrenc, D. T. Mason, and S. K. Ault. 1987. Exotic stock of Trypanosoma cruzi (Schizotrypanum) capable of development in and transmission by Triatoma protracta protracta from California: public health implications. Am. J. Trop. Med. Hyg. 36:523–528. Thurman, D. C. 1948. Key to Florida Triatoma with additional distribution records for the species (Hemiptera: Reduviidae). Florida Entomol. 31:58–62. Tomlinson, M. J., W. L. Chapman, Jr., W. L. Hanson, and H. S. Gosser. 1981. Occurrence of antibody to Trypanosoma cruzi in dogs in the southeastern United States. Am. J. Vet. Res. 42:1444–1446. Torres-Estrada, J. L., J. A. Martinez-Ibarra, and J. A. Garcia-Perez. 2002. Selection of resting sites of Triatoma gerstaeckeri (Stal) (Hemiptera: Reduviidae) females under laboratory and field conditions. Folia Entomol. Mexicana 41:63–66. Torrico, F., et al. 2004. Maternal Trypanosoma cruzi infection, pregnancy

290.

291.

292. 293.

294. 295. 296.

297. 298.

299.

300.

301.

302.

303.

304.

305.

306. 307.

308. 309.

310.

311.

312. 313. 314.

315. 316.

317. 318. 319.

679

outcome, morbidity, and mortality of congenitally infected and non-infected newborns in Bolivia. Am. J. Trop. Med. Hyg. 70:201–209. Traina, M. I., et al. 2010. Community-based study of Chagas disease prevalence in Los Angeles County, abstr. 497. Abstr. 59th Annu. Meet. Am. Soc. Trop. Med. Hyg., Atlanta, GA, 3 to 7 November 2010. Traina, M. I., et al. 2009. Prevalence of Chagas disease in U.S. Latin American immigrant population with cardiomyopathy, abstr. 408. Abstr. 58th Annu. Meet. Am. Soc. Trop. Med. Hyg., Washington, DC, 18 to 22 November 2009. Tyler, K. M., and D. M. Engman. 2001. The life cycle of Trypanosoma cruzi revisited. Int. J. Parasitol. 31:472–481. Umezawa, E. S., et al. 1999. Evaluation of recombinant antigens for serodiagnosis of Chagas’ disease in South and Central America. J. Clin. Microbiol. 37:1554–1560. Upton, S. J., R. A. Fridell, and M. Tilley. 1989. Trypanosoma kansasensis sp. n. from Neotoma floridana in Kansas. J. Wildl. Dis. 25:410–412. Urbina, J. A. 2009. New advances in the management of a long-neglected disease. Clin. Infect. Dis. 49:1685–1687. Usinger, R. L. 1944. The Triatominae of North and Central America and the West Indies and their public health significance. U.S. Government Printing Office, Washington, DC. Vaidian, A. K., L. M. Weiss, and H. B. Tanowitz. 2004. Chagas’ disease and AIDS. Kinetoplastid Biol. Dis. 3:2. Vakalis, N., J. H. Miller, E. Lauritsen, and D. Hansen. 1983. Anti-Trypanosoma cruzi antibodies among domestic dogs in New Orleans. J. Louisiana State Med. Soc. 135:14–15. Verani, J., et al. 2009. Geographic variation in the sensitivity of recombinant antigen-based rapid tests for chronic Trypanosoma cruzi infection. Am. J. Trop. Med. Hyg. 80:410–415. Verani, J. R., S. P. Montgomery, J. Schulkin, B. Anderson, and J. L. Jones. 2010. Survey of obstetrician-gynecologists in the United States about Chagas disease. Am. J. Trop. Med. Hyg. 83:891–895. Viotti, R., C. Vigliano, H. Armenti, and E. Segura. 1994. Treatment of chronic Chagas’ disease with benznidazole: clinical and serologic evolution of patients with long-term follow-up. Am. Heart J. 127:151–162. Viotti, R., et al. 2006. Long-term cardiac outcomes of treating chronic Chagas disease with benznidazole versus no treatment: a nonrandomized trial. Ann. Intern. Med. 144:724–734. Virreira, M., et al. 2006. Congenital Chagas disease in Bolivia is not associated with DNA polymorphism of Trypanosoma cruzi. Am. J. Trop. Med. Hyg. 75:871–879. Walsh, J. D., and J. P. Jones. 1962. Public health significance of the conenosed bug, Triatoma protracta (Uhler), in the Sierra Nevada foothills of California. Calif. Vector Views 9:33–37. Walton, B. C., P. M. Bauman, L. S. Diamond, and C. M. Herman. 1958. The isolation and identification of Trypanosoma cruzi from raccoons in Maryland. Am. J. Trop. Med. Hyg. 7:603–610. Wegner, D. H., and R. W. Rohwedder. 1972. The effect of nifurtimox in acute Chagas’ infection. Arzneimittelforschung 22:1624–1635. Wendel, S., and Z. Brener. 1992. Historical aspects, p. 5–12. In S. Wendel, Z. Brener, M. E. Camargo, and A. Rassi (ed.), Chagas disease-American trypanosomiasis: its impact on transfusion and clinical medicine. International Society of Blood Transfusion, Brazil, Sao Paulo, Brazil. Wendel, S., and D. A. Leiby. 2007. Parasitic infections in the blood supply: assessing and countering the threat. Dev. Biol. (Basel) 127:17–41. Westenberger, S. J., C. Barnabe, D. A. Campbell, and N. R. Sturm. 2005. Two hybridization events define the population structure of Trypanosoma cruzi. Genetics 171:527–543. Westenberger, S. J., N. R. Sturm, and D. A. Campbell. 2006. Trypanosoma cruzi 5S rRNA arrays define five groups and indicate the geographic origins of an ancestor of the heterozygous hybrids. Int. J. Parasitol. 36:337–346. WHO Expert Committee. 2002. Control of Chagas disease. WHO technical report series number 905. World Health Organization, Geneva, Switzerland. Williams, G. D., et al. 1977. Naturally occurring trypanosomiasis (Chagas’ disease) in dogs. J. Am. Vet. Med. Assoc. 171:171–177. Williams, J. T., E. J. Dick, Jr., J. L. VandeBerg, and G. B. Hubbard. 2009. Natural Chagas disease in four baboons. J. Med. Primatol 38:107–113. Wincker, P., et al. 1997. PCR-based diagnosis for Chagas’ disease in Bolivian children living in an active transmission area: comparison with conventional serological and parasitological diagnosis. Parasitology 114:367– 373. Wolf, A. F. 1969. Sensitivity to Triatoma bite. Ann. Allergy 27:271–273. Wood, F. 1934. Natural and experimental infection of Triatoma protracta Uhler and mammals in California with American human trypanosomiasis Am. J. Trop. Med. Hyg. 14:497–517. Wood, F. 1936. Trypanosoma neotomae, sp. nov., in the dusky-footed wood rat and the wood rat flea. Univ. Calif. Publ. Zool. 41:133–143. Wood, F. D., and S. F. Wood. 1937. Occurrence of haematozoa in some California birds and mammals. J. Parasitol. 23:197–201. Wood, S., and F. Wood. 1967. Ecological relationships of Triatoma p. pro-

680

320.

321.

322. 323.

324. 325. 326. 327. 328. 329. 330. 331.

332.

333.

334. 335. 336. 337.

CLIN. MICROBIOL. REV.

BERN ET AL. tracta (Uhler) in Griffith Park, Los Angeles, California. Pacific Insects 9:537–550. Wood, S. F. 1944. An additional California locality for Trypanosoma cruzi Chagas in the western cone-nosed bug, Triatoma protracta (Uhler). J. Parasitol. 30:199. Wood, S. F. 1949. Additional observations on Trypanosoma cruzi, Chagas, from Arizona in insects, rodents, and experimentally infected animals. Am. J. Trop. Med. 29:43–55. Wood, S. F. 1950. Allergic sensitivity to the saliva of the western cone-nosed bug. Bull. South. Calif. Acad. Sci. 49:71–73. Wood, S. F. 1962. Blood parasites of mammals of the California Sierra Nevada Foothills, with special reference to Trypanosoma cruzi and Hepatozoon leptosoma sp. n. Bull. South. Calif. Acad. Sci. 61:161–176. Wood, S. F. 1959. Body weight and blood meal size in conenose bugs, Triatoma and Paratriatoma. Bull. South. Calif. Acad. Sci. 58:116–118. Wood, S. F. 1941. Chagas’ disease (does it exist in men in Arizona?). Southwest Med. April 1941:112–114. Wood, S. F. 1965. Conenose bugs (Triatoma) visit unoccupied boy’s camp in Los Angeles. J. Med. Entomol. 1:347–348. Wood, S. F. 1951. Importance of feeding and defecation times of insect vectors in transmission of Chagas’ disease. J. Econ Entomol. 44:52–54. Wood, S. F. 1952. Mammal blood parasite records from Southwestern United States and Mexico. J. Parasitol. 38:85–86. Wood, S. F. 1975. New localities for mammal blood parasites from southwestern United States. J. Parasitol. 61:969–970. Wood, S. F. 1941. New localities for Trypanosoma cruzi Chagas in southwestern United States. Am. J. Trop. Med. Hyg. 34:1–13. Wood, S. F. 1975. Notes on possible natural control agents for conenose bugs: Triatoma and Paratriatoma (Hemiptera: Reduviidae). Natl. Pest Control Operator News 35:16–18. Wood, S. F. 1941. Notes on the distribution and habits of reduviid vectors of Chagas’ disease in the southwestern United States, part I. Pan-Pacific Entomol. 17:85–94. Wood, S. F. 1941. Notes on the distribution and habits of reduviid vectors of Chagas’ disease in the southwestern United States, part II. Pan-Pacific Entomol. 17:115–118. Wood, S. F. 1944. Notes on the feeding of the cone-nosed bugs (Hemiptera: Reduviidae). J. Parasitol. 30:197–198. Wood, S. F. 1942. Observations on vectors of Chagas’ disease in the United States. Calif. Bull. Soc. Calif. Acad. Sci. 41:61–69. Wood, S. F. 1943. Observations on vectors of Chagas’ disease in the United States. II. Arizona. Am. J. Trop. Med. Hyg. 23:315–320. Wood, S. F. 1946. The occurrence of Trypanosoma conorhini Donovan in

Caryn Bern is a medical epidemiologist in the Parasitic Diseases Branch, Division of Parasitic Diseases and Malaria, CDC. She holds an M.D. from Stanford University School of Medicine and an M.P.H. from the Johns Hopkins University School of Public Health and is board certified in internal medicine. She joined CDC as an Epidemic Intelligence Service Officer in 1990 and has worked in the Division of Parasitic Diseases since 1996. She holds adjunct appointments in the Emory Rollins School of Public Health and Johns Hopkins University School of Public Health and teaches tropical medicine courses at Johns Hopkins, the University of Minnesota, and the London School of Hygiene and Tropical Medicine. Dr. Bern’s current research is focused on the treatment, immunology, epidemiology, and control of Chagas’ disease and leishmaniasis.

338. 339. 340.

341.

342. 343.

344.

345. 346.

347. 348.

349.

350.

351. 352.

353.

the reduviid bug, Triatoma rubrofasciata (Degeer) from Oahu T. H. Proc. Haw. Entomol. Soc. 12:651. Wood, S. F. 1944. The reptile associates of wood rats and cone-nosed bugs. Bull. South. Calif. Acad. Sci. 43:44–48. Wood, S. F. 1975. Trypanosoma cruzi: new foci of enzootic Chagas’ disease in California. Exp. Parasitol. 38:153–160. Wood, S. F., and F. D. Wood. 1964. Nocturnal aggregation and invasion of homes in southern California by insect vectors of Chagas’ disease. J. Econ. Entomol. 57:775–776. Wood, S. F., and F. D. Wood. 1961. Observations on vectors of Chagas’ disease in the United States. III. New Mexico. Am. J. Trop. Med. Hyg. 10:155–162. Woody, N. C., and H. B. Woody. 1955. American trypanosomiasis (Chagas’ disease); first indigenous case in the United States. JAMA 159:676–677. World Health Organization. 2008. The global burden of disease: 2004 update. World Health Organization, Geneva, Switzerland. http://www.who .int/healthinfo/global_burden_disease/2004_report_update/en/index.html. World Health Organization. 2007, posting date. New global effort to eliminate Chagas disease (press release). http://www.who.int/mediacentre/news /releases/2007/pr36/en/index.html. Yabsley, M. J., and G. P. Noblet. 2002. Seroprevalence of Trypanosoma cruzi in raccoons from South Carolina and Georgia. J. Wildl. Dis. 38:75–83. Yabsley, M. J., G. P. Noblet, and O. J. Pung. 2001. Comparison of serological methods and blood culture for detection of Trypanosoma cruzi infection in raccoons (Procyon lotor). J. Parasitol. 87:1155–1159. Yaeger, R. G. 1961. The present status of Chagas’ disease in the United States. Bull. Tulane Univ. Med. Fac. 21:9–13. Yaeger, R. G. 1988. The prevalence of Trypanosoma cruzi infection in armadillos collected at a site near New Orleans, Louisiana. Am. J. Trop. Med. Hyg. 38:323–326. Yeo, M., et al. 2005. Origins of Chagas disease: Didelphis species are natural hosts of Trypanosoma cruzi I and armadillos hosts of Trypanosoma cruzi II, including hybrids. Int. J. Parasitol. 35:225–233. Yoshida, N. 2008. Trypanosoma cruzi infection by oral route: how the interplay between parasite and host components modulates infectivity. Parasitol. Int. 57:105–109. Young, C., P. Losikoff, A. Chawla, L. Glasser, and E. Forman. 2007. Transfusion-acquired Trypanosoma cruzi infection. Transfusion 47:540–544. Zeledon, R., R. Alvarado, and L. F. Jiron. 1977. Observations on the feeding and defecation patterns of three triatomine species (Hemiptera: Reduviidae). Acta Trop. 34:65–77. Zingales, B., et al. 2009. A new consensus for Trypanosoma cruzi intraspecific nomenclature: second revision meeting recommends TcI to TcVI. Mem. Inst. Oswaldo Cruz 104:1051–1054.

Sonia Kjos is currently a Project Scientist in the Epidemiology Center at the Marshfield Clinic Research Foundation in Marshfield, WI. She holds an M.S. in epidemiology from The University of Texas School of Public Health, Houston, and a Ph.D. in medical and veterinary entomology from Texas A&M University, College Station, where she also completed a certification program in geographic information systems. Dr. Kjos spent 2 years in the Division of Parasitic Diseases, CDC, Atlanta, GA, as an American Society for Microbiology postdoctoral fellow. Her research focus has been on the ecoepidemiology of peridomestic Chagas’ disease transmission in the United States, particularly the characteristics of the insect vectors and role of domestic animals as disease reservoirs.

VOL. 24, 2011

T. CRUZI AND CHAGAS’ DISEASE IN THE UNITED STATES

Michael J. Yabsley is currently an Associate Professor of Wildlife Disease Ecology at the University of Georgia. He has an M.S. in zoology (parasitology) from Clemson University and a Ph.D. in infectious diseases from the College of Veterinary Medicine at the University of Georgia. Dr. Yabsley teaches several courses in veterinary parasitology and principles of wildlife diseases at the undergraduate, graduate, and veterinary student levels and mentors undergraduate, M.S., and Ph.D. students interested in various aspects of wildlife disease ecology. He has an active research program that principally investigates the role of wildlife as reservoirs or hosts for zoonotic and/or vector-borne pathogens. Since 1997, Dr. Yabsley has been studying the natural history of T. cruzi in the United States. He is an active member of the American Society of Parasitologists, the Southeastern Society of Parasitologists, and the Wildlife Disease Association. He is the author or coauthor of 112 peer-reviewed publications.

681

Susan P. Montgomery is currently the Team Lead for the Epidemiology Team in the Parasitic Diseases Branch, Division of Parasitic Diseases and Malaria, CDC. She has a D.V.M. from New York State College of Veterinary Medicine, Cornell University, and an M.P.H. from the Harvard School of Public Health. After 15 years in private veterinary medical practice, she joined CDC as an Epidemic Intelligence Service Officer in 2002 with the Division of Vector-Borne Infectious Diseases and was a staff epidemiologist with the Foodborne and Diarrheal Diseases Branch before joining the Division of Parasitic Diseases in 2005. Her current research activities focus on Chagas’ disease epidemiology in the United States, schistosomiasis epidemiology and control, neglected infections associated with poverty in the United States, and parasitic disease blood and organ transplantation safety.

CLINICAL MICROBIOLOGY REVIEWS, Oct. 2011, p. 682–700 0893-8512/11/$12.00 doi:10.1128/CMR.00003-11 Copyright © 2011, American Society for Microbiology. All Rights Reserved.

Vol. 24, No. 4

Current Knowledge of Trichosporon spp. and Trichosporonosis Arnaldo L. Colombo,1* Ana Carolina B. Padovan,1 and Guilherme M. Chaves2 Laborato ´rio Especial de Micologia, Disciplina de Infectologia, Universidade Federal de Sa ˜o Paulo, Sa ˜o Paulo, Brazil,1 and Laborato ´rio de Micologia Me´dica e Molecular, Departamento de Ana ´lises Clínicas e Toxicolo ´gicas, Universidade Federal do Rio Grande do Norte, Natal, Brazil2 INTRODUCTION .......................................................................................................................................................682 CURRENT TAXONOMY OF TRICHOSPORON SPP. ..........................................................................................683 VIRULENCE FACTORS OF TRICHOSPORON SPP. AND SOURCES OF INVASIVE INFECTIONS ........684 Biofilms ....................................................................................................................................................................685 Enzymes....................................................................................................................................................................685 Cell Wall Components ...........................................................................................................................................685 Sources of Superficial and Invasive Infections...................................................................................................686 HUMAN INFECTIONS CAUSED BY MEMBERS OF THE GENUS TRICHOSPORON: AT-RISK POPULATIONS AND CLINICAL MANIFESTATIONS...............................................................................686 Superficial Infections..............................................................................................................................................687 Deep-Seated Infections...........................................................................................................................................687 DIAGNOSIS OF INVASIVE TRICHOSPORONOSIS AND LABORATORY TOOLS FOR IDENTIFICATION OF TRICHOSPORON SPP..............................................................................................688 Antigen Detection....................................................................................................................................................689 PCR-Based Methods...............................................................................................................................................689 Luminex xMAP Technology...................................................................................................................................690 New Perspectives on Microscopic Imaging Diagnosis .......................................................................................690 Phenotypic Identification of Trichosporon Species .............................................................................................691 Molecular Targets and Identification of Trichosporon Species in Cultures ...................................................692 PCR-based methods............................................................................................................................................692 Proteomics as a tool for fungal identification ................................................................................................693 Genotyping of Trichosporon spp., with an emphasis on the geographic distribution of T. asahii genotypes ..........................................................................................................................................................694 ANTIFUNGAL SUSCEPTIBILITY TESTING FOR TRICHOSPORON SPP. AND PRINCIPLES OF THERAPY ............................................................................................................................................................694 Antifungal Susceptibility Tests .............................................................................................................................694 Antifungal Therapy.................................................................................................................................................695 SUMMARY AND CONCLUSION............................................................................................................................695 ACKNOWLEDGMENTS ...........................................................................................................................................696 REFERENCES ............................................................................................................................................................696 The genus Trichosporon is phenotypically characterized by the ability to form blastoconidia, true mycelia, and, most importantly, arthroconidia, asexual propagules that disarticulate from true hyphae. The presence of multilamellar cell walls and dolipores with or without parenthesomes is an important characteristic of Trichosporon spp. (66, 69). Nevertheless, some species possess other morphological structures that can also be used to differentiate them, including appresoria, macroconidia, or other morphological structures and processes. Trichosporon spp. are able to utilize different carbohydrates and carbon sources and to degrade urea, but members of this genus are nonfermentative. Cell cultures grow on Sabouraud dextrose agar as yeast colonies with colors ranging from white to cream, typically displaying aspects such as cerebriform and radial surfaces (39). Colonies may become dry and membranous with age (101). The genus Trichosporon has a long and controversial history. It was first designated in 1865 by Beigel, who observed this microorganism causing a benign hair infection. The word Trichosporon is derived from the Greek and represents a combination of Trichos (hair) and sporon (spores). This nomencla-

INTRODUCTION Trichosporon spp. are basidiomycetous yeast-like anamorphic organisms (Basidiomycota, Hymenomycetes, Tremelloidae, Trichosporonales) that are widely distributed in nature and found predominantly in tropical and temperate areas. These organisms can be found in substrates such as soil, decomposing wood, air, rivers, lakes, seawater, cheese, scarab beetles, bird droppings, bats, pigeons, and cattle. In humans, these fungal species occasionally are part of the gastrointestinal and oral cavity microbiota and can transiently colonize the respiratory tract and skin (22, 60, 68, 76, 101, 108, 170, 186, 188, 189). Recently, Silvestre et al. found that 11% of their 1,004 healthy male volunteers were colonized by Trichosporon species on their normal perigenital skin (scrotal, perianal, and inguinal sites of the body) (165). * Corresponding author. Mailing address: Laborato ´rio Especial de Micologia, Disciplina de Infectologia, Universidade Federal de Sa˜o Paulo, Sa˜o Paulo-SP. Rua Botucatu, 740, CEP 04023-062 Sa˜o Paulo, Brazil. Phone and fax: (55) (11) 5083-0806. E-mail: colomboal@terra .com.br. 682

VOL. 24, 2011

ture is related to the presence of irregular nodules along the body and head hair, consisting of a fungal infection, which is further characterized as white piedra (meaning “stone” in Spanish). Indeed, the etiological agent of this white piedra was formerly misclassified as the alga Pleurococcus beigelii (66, 69). In 1890, Behrend described the agent causing white piedra found on a man’s beard in detail and named it Trichosporon ovoides (16). Since then, other Trichosporon species have been reported. In 1902, Vuillemin designated all Trichosporon species Trichosporon beigelii, an arthrospore-containing yeast (66, 69). In 1909, Beurmann et al. cultured fungal elements collected from a patient with a pruritic cutaneous lesion and designated the isolated fungus Oidium cutaneum, which was subsequently renamed Trichosporon cutaneum by Ota in 1926. Nevertheless, in 1942 Diddens and Lodder considered T. beigelii and T. cutaneum to be the same species. This led to the use of two names for this fungus with clinical relevance: Trichosporon beigelii, adopted by physicians, and Trichosporon cutaneum, preferred by environmental mycologists (66, 69). Although most Trichosporon spp. routinely isolated in laboratories are related to episodes of colonization or superficial infections, this fungus has been recognized as an emergent opportunistic agent causing invasive infections in tertiary care hospitals worldwide (25). The genus Blastoschizomyces, which contains a unique species named B. capitatus, was also considered to belong to the Trichosporon genus in the past. It has been recognized as a rare but emerging cause of disseminated infection specifically in leukemic patients (32). B. capitatus was thereafter reassigned to the genus Geotrichum (Geotrichum capitatus), but since 1985 the species has been recognized as more closely related to the Ascomycetes despite the fact it produces blastoconidia and arthroconidia just like Trichosporon (157). Indeed, B. capitatus has a well-described teleomorph named Dipodascus capitatus which produces asci and ascospores (38). In addition, it produces annelloconidia, a distinct cell wall composition, and septal pores, peculiar characteristics that make it different from the genera Trichosporon and Geotrichum (157).

CURRENT TAXONOMY OF TRICHOSPORON SPP. Traditional taxonomy based on phenotypic characterization of fungal organisms generates inconsistent data when applied to the genus Trichosporon, as it aggregates within the same taxon isolates with genetically heterogeneous behavior. In 1982, some strains belonging to the taxon T. beigelii were found to possess a Q-10 ubiquinone system, while others contained a Q-9 system (192). Since then, several authors have reported clear differences among T. beigelii isolates based on physiological, morphological, and genetic characteristics. In 1991, Kemker et al. started to describe clearly the diversity among Trichosporon spp. isolated from environmental and clinical sources (94). These authors evaluated different isoenzymes present in 15 Trichosporon strains and the genetic profiles of these strains by using restriction fragment length polymorphisms (RFLP) analysis of ribosomal DNA (rDNA), including both the internal transcribed spacer 1 (ITS1) and ITS2 regions and the 5.8S gene as targets. These data were

TRICHOSPORON SPP. AND TRICHOSPORONOSIS

683

compared with morphological features, carbon substrate usage profiles, and uric acid assimilation abilities. In 1992, Gueho and collaborators conducted molecular studies of nucleic acids correlated with the morphological, physiological and biochemical characteristics of a collection of strains and proposed the reassignment of the genus Trichosporon (69). The criteria used to reclassify Trichosporon species were based on several aspects, including the ultrastructure of septal pores. These authors demonstrated that although all Trichosporon species contain an electron-dense and multilamellar cell wall, at least two basic types of dolipores exist. The first is composed of nailhead-like swellings separated by a narrow pore canal that is often filled by an electron-dense mass, whereas the second is constituted by well-defined, rounded inflations with a sponge-like or vesicular substructure covered by a parenthesome. Other characteristics, such as coenzyme Q systems Q-10 and Q-9, differed within Trichosporon spp. and corresponded to the guanine-plus-cytosine content (moles percent G⫹C). DNA reassociation values, nutritional profiles, and evaluation of part of the 28S regions of rDNA sequences also demonstrated differences between Trichosporon species. Following analysis of 101 strains, these authors described a new species (Trichosporon mucoides) and 19 taxa; additionally, some previously described species were designated synonyms of other species. In the same year, Gueho et al. reported on the variety of the genus and suggested that various series of species can be recognized based upon different niches (66). Strains differ based on their source of isolation (soil, water, or human or animal disease) and based on their phenotypic and molecular properties. The two species T. beigelii and T. cutaneum, considered to be synonyms since 1942, were clearly different as determined by ubiquinone systems and percent G⫹C DNA content. Therefore, the authors considered the nomenclature T. beigelii to be inappropriate and suggested that it be abandoned. In 1994, those authors concluded that the genus Trichosporon should include the following six species: T. cutaneum, T. asahii, T. asteroides, T. mucoides, T. inkin and T. ovoides (67). Therefore, the previous taxon T. beigelii was replaced by several species, and the taxonomy of the genus was progressively modified by powerful molecular tools able to discriminate between phylogenetically closely related species (66, 176, 177). In 1994 and 1995, Sugita et al. reviewed the genus Trichosporon and proposed a new classification including 17 species and 5 varieties of Trichosporon (176, 177). In 2002, Sugita et al. proposed 25 species for the genus Trichosporon and suggested that 8 should be considered relevant as potential human pathogens, including the two emergent species T. domesticum and T. montevideense (172). Thereafter, the same group published a report in 2004 recognizing 36 Trichosporon species, including 5 new species proposed by Middlehoven et al. in 2004: T. vadense, T. smithiae, T. dehoogii, T. scarabaeorum, and T. gamsii (169). In 2004, Middlehoven et al. also separated the order Trichosporonales into four clades named Gracile, Porosum, Cutaneum, and Ovoides (Table 1) (123). In that same year, Sugita et al. (169) included the clade Brassicae in the order Trichosporonales, which encompassed some species considered to belong to the Gracile clade by Middlehoven et al. (123) (Table 1). A new Trichosporon species isolated from the hindgut of the

684

COLOMBO ET AL.

CLIN. MICROBIOL. REV.

TABLE 1. Currently accepted Trichosporon species and their subdivision within different clades Species namea

Clade

Species no.

Gracile/Brassicae

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17

Trichosporon brassicae Trichosporon domesticum Trichosporon montevideense Trichosporon scarabaeorum Trichosporon mycotoxinivorans Trichosporon dulcitum Trichosporon cacaoliposimilis Trichosporon gracile Trichosporon laibachii Trichosporon multisporum Trichosporon vadense Trichosporon veenhuisii Trichosporon akiyoshidainum Trichosporon chiropterorum Trichosporon siamense Trichosporon otae Trichosporon loubieri

Cutaneum

18 19 20 21 22 23 24 25 26 27 28 29

Trichosporon cutaneum Trichosporon debeurmannianum Trichosporon dermatis Trichosporon jirovecii Trichosporon oleaginosus Trichosporon moniliiforme Trichosporon mucoides Trichosporon smithiae Trichosporon terricola Trichosporon middelhovenii Trichosporon shinodae Trichosporon cavernicola

Porosum

30 31 32 33 34 35 36 37 38 39 40 41 42 43 44

Trichosporon aquatile Trichosporon asahii Trichosporon asteroides Trichosporon caseorum Trichosporon coremiiforme Trichosporon faecale Trichosporon inkin Trichosporon japonicum Trichosporon lactis Trichosporon ovoides Trichosporon insectorum Trichosporon porosum Trichosporon dohaense Trichosporon chiarellib Trichosporon xylopini

Ovoides

45 46 47 48 49 50

Trichosporon Trichosporon Trichosporon Trichosporon Trichosporon Trichosporon

dehoogii gamsii guehoae lignicola sporotrichoides wieringae

a

The species in bold were described as being of medical interest. b Closely related but placed in a different clade by Pagnocca et al. (139).

lower termite Mastotermes darwiniensis, recognized as an important detoxifier of mycotoxins, was described by Molnar et al. in 2004 and named T. mycotoxinivorans, belonging to the clade Gracile (125). In 2005, a study addressing the isolation and identification of yeasts from guano of bat-inhabited caves in Japan proposed seven novel potential Trichosporon species. The new species were not representative of a single clade (Gracile, Porosum,

Brassicae, Cutaneum, or Ovoides) but were distributed across all but the Ovoides clade (170). In 2008, Fuentefria proposed a new Trichosporon species isolated from the guts of insects from Panama and from artesian cheese prepared in Brazil. The new species was included in the Ovoides clade (60). In 2009, a new Trichosporon species was described in Qatar. Taj-Aldeen et al. described a novel Trichosporon species named T. dohaense following the investigation of 27 clinical isolates of Trichosporon obtained from immunosuppressed patients. Three isolates of the new species were cultured from patients with onychomycosis, catheter-related infection, and tinea pedis. The novel species is closely related to T. coremiiforme and belongs to the Ovoides clade (180). In 2010, in Brazil, a new Trichosporon species was described based on the analysis of the D1/D2 and ITS regions of 34 Trichosporon strains closely related to T. scarabaeorum. The new species was named T. chiarelli, and strains were isolated from a fungus garden and from waste. This novel species was considered a sibling species of the Cutaneum clade (139). Recently, other Trichosporon species have been described. The first species was originally cultured from the gut of the wood-resident beetle Xylopinus saperdioides and named T. xylopini. It is clearly distinguishable from its closest relative, T. porosum, by 14 nucleotides in the ITS and D1/D2 regions of the rRNA genes. Of note, T. xylopini has been recognized as an arthroconidium-producing yeast that is able to degrade hemicellulose (71). Two other novel Trichosporon species were characterized based on the analysis of oleaginous yeasts maintained at the American Type Culture Collection (ATCC Mycology Collection). They were named T. cacaoliposimilis and T. oleaginosus, respectively. Molecular phylogenetic analyses showed that T. cacaoliposimilis belongs to the Gracile clade, close to T. gracile and T. dulcitum, whereas T. oleaginosus belongs to the Cutaneum clade, with T. jirovecii as the closest taxon (70). In conclusion, as illustrated in Table 1, 50 species of Trichosporon have been described from different regions of the globe, including 16 species with clinical relevance (139). The previously recognized taxon T. pullulans, which was considered to belong to the genus Trichosporon by Diddens and Lodder in 1942, has now been reassigned to a new genus and is named Guehomyces pullulans (51). Based on the large variability of natural habitats related to all different species of Trichosporon, it is possible to suggest that different reservoirs may play a role in human infections. VIRULENCE FACTORS OF TRICHOSPORON SPP. AND SOURCES OF INVASIVE INFECTIONS Fungi that are opportunistic pathogens retain several factors that allow their growth and permit the establishment of disease and their dissemination within the host. These factors are known as virulence factors. Such factors are usually related to morphological switching, the ability to adhere to abiotic surfaces, thermotolerance, the expression of cell wall components, and enzyme production and secretion (77, 87, 145). Even though Trichosporon spp. can represent the second or third most common non-Candida yeast infections causing invasive disease in patients with hematological cancer, few reports have addressed the virulence factors of this genus (24, 42, 138).

VOL. 24, 2011

TRICHOSPORON SPP. AND TRICHOSPORONOSIS

Biofilms Infections with invasive Trichosporon spp. are usually associated with central venous catheters, vesical catheters, and peritoneal catheter-related devices. The ability to adhere to and form biofilms on implanted devices can account for the progress of invasive trichosporonosis, as it can promote the escape from antifungal drugs and host immune responses. Di Bonaventura et al. were the first to use electron microscopy to analyze the kinetics of biofilm formation and development on polystyrene surfaces. They tested 4 T. asahii strains, 3 of which were isolated from blood samples of patients with hematological malignancies (42). Those authors showed that, similarly to data described for biofilms of Candida spp., T. asahii cells were able to rapidly adhere to polystyrene after a 30-min incubation. These cells presented different morphologies, such as budding yeasts and filamentous forms, embedded within an extracellular polysaccharide (EPS) matrix composing the ultrastructure of the mature biofilm after 72 h. Analysis of confocal imaging sections taken across the biofilm showed that the thickness of mature biofilms varied from 25 to 40 ␮m. These authors also tested the susceptibility profiles of planktonic and biofilm cells to amphotericin B, caspofungin, fluconazole, and voriconazole. T. asahii biofilms were resistant to all antifungals tested (MIC ⬎ 1,024 ␮g/ml) and were up to 16,000 times more resistant to voriconazole than planktonic cells (MIC ⫽ 0.06 ␮g/ml). Enzymes Another important virulence factor is the ability to produce and secrete enzymes for scavenging nutrients from the environment. Proteases and phospholipases are among the enzymes that can increase fungal pathogenicity by breaking up proteins and disrupting host cell membranes, depending on their expression levels and the host immune response (61). In 1994, Chen and collaborators purified two lipase enzymes (lipases I and II) from T. fermentans WU-C12 and verified that these enzymes could hydrolyze olive oil with an optimum pH of 5.5 and temperature of 35°C (31). These lipases also demonstrated stable activity after incubation at 30°C for 24 h over a pH range of 4.0 to 8.0 and were further characterized by cDNA isolation and sequencing (7). BLAST (Basic Local Alignment Search Tool, http://blast.ncbi.nlm.nih.gov/) analysis showed that the cDNA encoded by the gene TFL1 and the putative protein sequence had 99.5% similarity to those of lipase II from Geotrichum candidum. Southern blot analysis revealed that there were two lipase genes carried on the genome, confirming the prior purification of two lipases. A putative correlation between the ability to produce Tfl1 lipase and the virulence of Trichosporon in humans has not been examined. However, one can speculate that lipases in general are important for the invasion process and for adaptation to different tissues within the host. With regard to Trichosporon spp. isolated from infected patients, Chaves and collaborators have analyzed the proteinase and phospholipase activities of a collection of 30 yeast isolates, including two strains of T. pullulans (30). Both of the T. pullulans isolates did not exhibit phospholipase activity but did display proteinase activity. This result is reasonable in light of

685

the reclassification of T. pullulans as Guehomyces pullulans, which could be a phospholipase nonproducer. Dag and Cerikc¸ioglu analyzed the production of a variety of virulence factors from 50 T. asahii strains, the majority of which were isolated from urine, although some originated each from blood, peritoneal fluid, nephrostomy, tongue swab, and nail. Proteinase and phospholipase activities could not be detected from these strains, although all isolates were esterase positive. Slime production was moderate in 10 strains, while 18 strains were characterized as weak producers and 20 strains were defined as slime nonproducers (37). Later, Cafarchia and collaborators investigated the phospholipase activity in a collection of 163 yeast isolates recovered from pigeon cloacae and droppings (22). Biochemical and morphological tests identified 13 species among these isolates, of which 14 isolates were identified as T. beigelii. Two of the 10 isolates recovered from cloacae exhibited phospholipase activity in egg yolk medium plates after 2 and 5 days of incubation, whereas all 4 isolates from excreta exhibited phospholipase activity. This was the first report showing phospholipase activity in Trichosporon spp., and no gene in the genome has been described to have this function. Recently, an alkaline lipase was isolated from T. asahii MSR 54, which was previously cultured from petroleum sludge in India for application in developing a presoak formulation for oil removal (100). The enzyme was active at ambient temperature but presented maximal activity at 40°C and over a pH range of 8.0 to 10.0. An analysis of the production of extracellular enzymes and the morphology switch of 61 clinical isolates of T. asahii revealed that these isolates were not able to produce secreted aspartic proteinases or phospholipases, while they were able to secrete active beta-N-acetylhexosaminidase. No other clinically relevant Trichosporon spp. have been shown to produce this enzyme (84). The same study also reported 4 different morphological types of colonies grown on Sabouraud dextrose agar (SDA), i.e., white farinose (69%), white pustular (18%), yellowish white (10%), and white cerebriform (3%). Those authors observed that strains of the three major types usually developed two to five colony types and switched at a frequency of 102 to 104 when grown at 37°C, similar to that observed with Candida albicans and Cryptococcus neoformans (4, 59, 87). Most of the colonies switched irreversibly to the smooth type, which had the greatest beta-N-acetylhexosaminidase enzymatic activity compared with the parent type in all strains (84). Despite the available information on the capability of Trichosporon strains to produce lipases and proteases, the roles of these enzymes in the pathogenesis of human invasive infections are not clear. In addition, this genus may possess cryptic enzymes not yet described. Cell Wall Components Members of Trichosporon spp. express glucuronoxylomannan (GXM) in their cell walls, similarly to C. neoformans. GXM is a 1,3-linked mannan backbone attached to short side chains of 1,4-linked mannose and 1,2-linked xylose residues by substituting the 2 or 4 portion of the 1,3-linked mannose residues of the main group (124). This polysaccharide may attenuate the phagocytic capability of neutrophils and monocytes in

686

COLOMBO ET AL.

vivo (150). Karashima and collaborators reported that T. asahii isolates could switch phenotype and increase the amount of secreted GXM after passage in a mouse model (91). Three environmental isolates were analyzed before and after consecutive passages through mice and compared with 14 isolates from patients with deep-seated infections or recovered from lung autopsies, blood, urine, sputum, and catheters. That study demonstrated that the colonies of all of the environmental isolates were rugose, whereas the majority of the clinical isolates were powdery when grown on SDA. The clinical isolates consisted of 90% blastoconidia and arthroconidia, whereas all of the environmental isolates consisted of 99% hyphae. Interestingly, all environmental isolates from mouse kidneys recovered in SDA switched their macromorphologies to powdery colonies, with conidia as the predominant cell type. All clinical isolates released significantly higher levels of GXM than did environmental isolates (titers of log2 9.4 ⫾ 0.7 versus log2 5.4 ⫾ 1.4). After passage in mice, environmental isolates generated increased GXM concentrations (titers of log2 10.0 ⫾ 0.7 versus log2 5.4 ⫾ 1.4). Minor changes in the (133)-␤-D-glucan (BDG) titers were observed, and no significant difference in titers was detected for environmental isolates before and after passage in mice. Taken together, the changes in the phenotype of the environmental isolates after passages in mice seem to be a regular response to adaptation within the host in order to facilitate dissemination and escape from phagocytosis by polymorphonuclear leukocytes and monocytes in vivo. The protective role of GXM was also addressed by analyzing the structure and function of GXM isolated from T. asahii and testing whether this polysaccharide could protect acapsular C. neoformans mutants from phagocytosis by mouse macrophages. Fonseca and collaborators reported that the concentration of GXM in the supernatants of T. asahii cultures was approximately 12 times less than that in C. neoformans supernatants (56). The major components of T. asahii GXM were mannose (60%), followed by xylose (24%) and glucose (8%). This differed from the case for C. neoformans supernatants, from which the third most commonly isolated component was glucuronic acid (10%). Fluorescence microscopy also demonstrated that T. asahii cells possessed less GXM in their cell walls than C. neoformans cells. To test the protective effect of GXM against phagocytosis, acapsular C. neoformans mutant cells were coated with GXM purified from a C. neoformans high-GXM producer and with GXM from T. asahii and incubated with murine macrophage cells in vitro. Yeast cells containing no detectable surface GXM were rapidly internalized by phagocytes, and the polysaccharide-coated cells were phagocytized 6 times less frequently. However, no significant difference was observed with regard to the GXM source, indicating that despite differences in major sugar components, GXM from T. asahii had a protective role similar to that of GXM from C. neoformans (56, 84). Sources of Superficial and Invasive Infections The source of superficial and deep-seated Trichosporon infections is still the subject of considerable debate. It is important to consider that Trichosporon spp. are widely distributed in nature and may be part of the normal biota of the skin, respiratory tract, gastrointestinal tract, and vagina (24, 187).

CLIN. MICROBIOL. REV.

The mode of transmission of superficial trichosporonosis remains unclear, but poor hygiene habits, bathing in contaminated water, and sexual transmission may play a role (17, 96). Although there is no consensus on the route of infection of human beings, close contact with an incident case, hair humidity, and the length of the scalp hair have all been recognized as risk factors for acquiring white piedra (96, 153, 158, 197). In regard to deep-seated infections, gastrointestinal colonization and further translocation throughout the gut may be considered the source of infection in a considerable number of episodes of trichosporonosis documented in cancer patients. In support of an endogenously acquired route of infection, we may look to evidence in support of gut translocation as the major source of candidemia, a biological model that may explain invasive infections caused by yeast pathogens that colonize the gastrointestinal tract (34, 134). In developing the first animal model of invasive trichosporonosis, Walsh and colleagues showed that Trichosporon was able to disseminate from the gut to the blood and other organs in rabbits after immunosuppression but not in healthy animals (189). In addition to endogenous acquisition from the gut, evidence also suggests that a substantial number of trichosporonosis cases may be exogenously acquired. Trichosporon spp. may enter the blood after contamination of a percutaneously inserted intravascular catheter via colonized skin. Indeed, Kontoyiannis et al. evaluated 17 cancer patients with trichosporonosis and suggested that central venous catheterrelated fungemia was the cause of 70% of all episodes (98). Interestingly, those authors suggested that prophylaxis with fluconazole, frequently used in patients with hematological disease, may prevent the gut translocation of Trichosporon spp. in this population, making the exogenous acquisition of this pathogen (i.e., mediated by colonization of a central venous catheter) the most plausible source of infection in this specific population. Finally, infants with very low birth weight (VLBW) may develop trichosporonosis, despite it being a rare cause of sepsis in neonates. Either yeast gastrointestinal translocation or catheter-associated fungemia may occur in these patients. Neonate skin colonization by Trichosporon may be acquired either from health care workers or after vaginal delivery, as up to 14% of woman may harbor this organism in their vulvovaginal region (45, 156).

HUMAN INFECTIONS CAUSED BY MEMBERS OF THE GENUS TRICHOSPORON: AT-RISK POPULATIONS AND CLINICAL MANIFESTATIONS Trichosporon is a medically important genus whose members are able to colonize and proliferate in different parts of human body, including the gastrointestinal system, respiratory tract, skin, and vagina. This yeast-like pathogen may cause deepseated, mucosa-associated, or superficial infections. Invasive trichosporonosis is documented mostly in patients with hematological malignancies and other medical conditions associated with immunosuppression, whereas superficial infections and allergic pneumonia are found predominantly in immunocompetent hosts (24, 46, 98, 184). Table 2 summarizes the sources of infection, main associated conditions, and etiological agents

VOL. 24, 2011

TRICHOSPORON SPP. AND TRICHOSPORONOSIS

687

TABLE 2. Human trichosporonosis: sources of infection, main associated conditions, and etiological agents Infection category

Main type(s) of infection

Major agent(s)

Main associated conditions

Invasive

Fungemia, urinary tract infections, peritonitis, endocarditis, othersa

T. asahii, T. mucoides, T. asteroides

Allergic pneumonia

Summer-type hypersensitivity pneumonitis White piedra

T. cutaneumb

Cancer, vascular and urinary catheters, organ transplantation, broad-spectrum antibiotic therapy Hot and humid weather, environmental contamination Young age and female sex, long hair, humidity, poor hygiene, headband use

Superficial a b

T. inkin, T. cutaneum, T. ovoides, T. loubieri

Arthritis, esophagitis, meningitis, brain abscess, splenic abscess, and uterine infections. T. cutaneum is now considered a complex of 10 different species.

that are involved with the most clinically relevant Trichosporon infections. Superficial Infections The clinical presentation of Trichosporon infection in humans is often benign superficial lesions of hair, called white piedra, characterized by the presence of irregular nodules on the affected hair. These nodules are loosely attached to the hair shaft, have a soft texture, and may be white or light brown. White piedra is a cosmopolitan disease affecting children and adults from areas with tropical and temperate climates. Most cases of white piedra have been reported in children and young adults, particularly females who frequently use headbands. Although rarely reported in the United States, this disease may be found in Texas and may be underreported in several regions (63, 96, 159). White piedra may be found in a large variety of hairy regions, including the scalp, beard, moustache, eyebrows, axilla, and, particularly, genital hairs. Concomitant hair infection by Trichosporon spp. and corynebacteria has been demonstrated by electron microscopy and histochemical evaluation of concretions, and it is still unclear whether the proteolytic capabilities of the coryneform bacteria may be relevant for the establishment of fungal infection (53, 54, 65, 67, 92, 96, 177, 197). White piedra appears to be caused predominantly by T. inkin, T cutaneum, T. ovoides, and T. loubieri, which has recently been reported in the literature as an emergent species related mainly to superficial infections in humans (9, 67, 99, 137, 177). It is noteworthy that Trichosporon spp. can also cause other superficial infections, such as onychomycosis, where the more frequently isolated species is T. cutaneum. Indeed, some Mexican authors have documented that the isolation of Trichosporon spp. from tinea pedis and onychomycosis patients may range from 2.81% to 42.8% of cases (9, 121, 155). Deep-Seated Infections After the first case of invasive trichosporonosis described by Watson and Kallichurum in 1970 (190a), several cases of invasive trichosporonosis in different clinical scenarios were described. For instance, in 1982 Manzella et al. reported a case of fungemia with cutaneous dissemination due to a Trichosporon sp. documented in a leukemic patient followed until death (113). Some years later, in 1988, Reinhart et al. reported a case of endocarditis caused by a Trichosporon sp. in a patient who

had previously suffered from rheumatic disease (148). In 1997, Lopes et al. described a case of peritonitis due to T. inkin in a diabetic patient (107). In 2001, Moretti-Branchini et al. reported 2 cases of invasive Trichosporon infections involving 2 patients with bone marrow transplants hospitalized at the Clinical Hospital of The University of Campinas, Brazil (126). A case of chronic dissemination infection due to a Trichosporon sp. resulting in multiple liver abscesses was reported by Meyer et al. in 2002 (122). In 2005, Abdala et al. reported a case of invasive T. asahii infection in a nonneutropenic patient undergoing orthotopic liver transplantation who died despite treatment with amphotericin B (1). After combining all of the reported cases of trichosporonosis, one can suggest that Trichosporon spp. appear to be increasing agents of invasive mycoses in contemporary medicine. In patients with malignant hematological diseases, this genus has been reported as the second most common agent of disseminated yeast infections, behind only the genus Candida, leading to 50 to 80% mortality rates despite treatment with antifungal therapy (55, 99, 138, 188). Indeed, breakthrough trichosporonosis in immunocompromised patients after administration of amphotericin B and echinocandins, and more rarely after the use of triazoles, has been extensively reported (15, 64, 79, 118, 186). The largest retrospective multicenter study of invasive trichosporonosis and geotrichosis in patients with malignant hematological diseases was conducted by Girmenia et al. in 2005 and included data on Trichosporon and Geotrichum infections documented over 20 years (62). Those authors reviewed 287 cases of trichosporonosis and 99 cases of geotrichosis documented from all over the world. The most common underlying conditions related to trichosporonosis were hematological diseases, peritoneal dialysis, and solid tumors. Trichosporonemia occurred in 115/154 (74.7%) of patients and disseminated infection in 78/154 (50.6%) of cases. The majority of the cases of trichosporonosis and geotrichosis were reported in North American medical centers (33.9%), followed by medical centers from Europe (27.6%) and Asia (23.3%). Only 6 isolates from South American institutions were reported, including 5 Brazilian isolates and 1 isolate from Argentina. Despite the large number of Trichosporon isolates included in this review, two were accurately identified to the species level as T. loubieri and 8 were assigned to the old taxon T. pullulans (recently named G. pullulans), while the other 287 isolates were identified only as Trichosporon sp. In 2004, Kontoyannis et al. described the clinical spectrum and outcome of 17 patients with cancer and invasive tricho-

688

COLOMBO ET AL.

sporonosis documented at the MD Anderson Cancer Center. The overall incidence of invasive trichosporonosis was found to be 8 cases per 100,000 admitted patients; 65% of the infected patients had acute leukemia, and 65% had neutropenia. Most patients (59%) had fungemia as the sole manifestation of the fungal infection, and 7 of 10 with Trichosporon fungemia had a central venous catheter-related infection. Of note, 60% of episodes were documented in patients who had been exposed to at least 7 days of antifungal therapy (breakthrough infections). The crude mortality rate at 30 days after admission was 53% (98). In 2009, Ruan et al. described a series of 19 patients with invasive trichosporonosis documented between 2000 and 2008 at the National Taiwan University Hospital. Cancer was the underlying disease in 58% of patients, and only 4 patients (21%) were neutropenic at the time of the diagnosis. Central venous catheter placement and the use of antibiotics were the most commonly associated conditions, being present in 90% and 95% of all patients, respectively. The crude mortality rate at 30 days after infection was 42% (154). In 2010, Suzuki et al. retrospectively evaluated clinical aspects and outcomes for 33 patients with Trichosporon fungemia and hematological malignancies in 5 different Japanese tertiary care centers between 1992 and 2007. The majority of these patients had acute leukemia (82%) and neutropenia (85%), and 90% of them had been exposed to at least 5 days of systemic antifungal therapy (breakthrough infections). Skin lesions were reported in 12 patients and pneumonia in 19 patients. The mortality rate attributable to the fungus was found to be 76%, with 67% of deaths occurring within 10 days of admission (179). Critically ill patients admitted to intensive care units and subjected to invasive medical procedures and antibiotic treatment are the patients most commonly associated with trichosporonosis, other than patients with malignant diseases (24, 25, 44, 154, 191). A recent paper from Chagas-Neto et al. (25) showed that in a series of 22 Brazilian pediatric and adult patients with trichosporonosis fungemia, most were ICU patients with no malignant disease. Most infected patients had degenerative diseases with organ failures and were subjected to treatment with broad-spectrum antibiotics and multiple invasive medical procedures before developing fungemia. Crude mortality rates were 30% for children and 87.5% for adult patients (P ⫽ 0.025) (24, 25). Finally, small series of cases of invasive trichosporonosis have also been described for burn victims, patients with secondary hemochromathosis, and Job’s syndrome (23, 26, 73, 162). Based on these series of cases, it is possible that fungemia and fever represent the most common clinical findings described for Trichosporon hematogenic dissemination. However, despite being less frequent, there are increasing reports of episodes of organ-specific infections and patients with disseminated trichosporonosis presenting with pneumonia, soft tissue lesions and eventually endophthalmitis, endocarditis, brain abscess, meningitis, arthritis, esophagitis, lymphadenopathy, liver, splenic abscess, and uterine infections (23, 24, 26, 27, 74, 98, 110, 117, 154, 178). Peritonitis is one of the most common and serious complications of peritoneal dialysis, and a case of peritoneal infection due to Trichosporon spp. in this setting was documented by

CLIN. MICROBIOL. REV.

Khanna et al. in 1980 (95). Fungal peritonitis is usually documented in patients with a history of previous bacterial infections, and this condition presents with sign and symptoms usually associated with any microorganism causing dialysisassociated peritonitis: fever, abdominal fullness, and turbid peritoneal dialysate fluid (88, 111). Despite being considered rare, endocarditis due to Trichosporon species in native or prosthetic valves of nonneutropenic patients has been increasingly reported. Patients usually develop large vegetations that may lead to embolic phenomena frequently located on the aortic bifurcation, in arteries of the lower extremities, or eventually in the brain. As suggested by most reports, valve replacement is mandatory, but recurrence of infection is very common and prognosis is generally poor regardless of the antifungal therapy (29, 86, 93, 114, 116, 147– 149, 164, 182). Urinary tract infections by this pathogen may also occur, especially in patients with urinary obstruction or those undergoing vesical catheterization and antibiotic treatment. These infections represent a clinical challenge for clinicians, as there are no clear and specific indications for the clinical interpretation of Trichosporon sp. recovery in urine. Although unusual, renal damage and aggravation of renal dysfunction may occur (50, 127, 191). Although most published reports on Trichosporon disseminated infections present scarce data in terms of species identification of the etiological agents using molecular methods, T. asahii is the most frequently isolated species in these infections (25, 154, 179). Finally, Ando et al. have described that the genus Trichosporon, in addition to causing infections, is the major etiological agent of summer-type hypersensitivity pneumonitis (SHP) in Japan (5). The authors investigated 621 patients with SHP and found that 95% of them had anti-Trichosporon antibodies in serum. Trichosporon isolates were found extensively in patients’ houses, and the elimination of these organisms from patients’ houses prevented disease (6, 163, 195, 196). Trichosporon species are responsible for SHP, leading to type III and IV allergies via repeated inhalation of organic dust and fungal arthroconidia that contaminate home environments during the hot, humid, and rainy summer season in western and southern Japan (11, 65, 92, 133, 169, 181, 188, 194). SHP is an immunologically induced lung disease, with pathogenesis mechanisms involving an initial immune complex-mediated lung injury followed by cell-mediated tissue damage with massive lymphocyte infiltration into the lungs (90). Mizobe et al. have characterized the antigenic components involved in SHP as GXM (124). T. cutaneum was originally described as the major agent of SHP, exhibiting 4 different genotypes: I, II, III, and I-III. After the extensive revision of the taxonomy of Trichosporon spp., the taxon T. cutaneum was divided into more than 10 different species, which tend to be grouped within the previously described serotypes (6, 85, 133, 169). DIAGNOSIS OF INVASIVE TRICHOSPORONOSIS AND LABORATORY TOOLS FOR IDENTIFICATION OF TRICHOSPORON SPP. In accordance with the definitions of opportunistic invasive fungal infections (IFI) published by the European Organization for Research and Treatment of Cancer/Invasive Fungal

VOL. 24, 2011

TRICHOSPORON SPP. AND TRICHOSPORONOSIS

Infection Cooperative Group (EORTC/IFICG) and the National Institute of Allergy and Infectious Disease Mycoses Study Group (NIAID/MSG), invasive trichosporonosis may be defined as follows (40). 1. “Proven invasive trichosporonosis.” Patients with proven invasive trichosporonosis presenting at least one of the following criteria: (i) blood cultures yielding Trichosporon species in patients with temporally related clinical signs and symptoms of infection, (ii) cerebrospinal fluid (CSF) cultures yielding Trichosporon species, or (iii) biopsy specimens that are culture positive and present histopathological evidence of fungal elements compatible with Trichosporon spp. 2. “Probable invasive trichosporonosis.” Patients with probable invasive trichosporonosis present all of the following criteria: (i) presence of at least one host factor (therapy with an immunosuppressive drug[s], neutropenia, or persisting fever despite therapy with appropriate broad-spectrum antibiotics), (ii) one microbiological criterion (culture or presence of fungal elements compatible with Trichosporon in a suspect biological material), and (iii) one major clinical criterion (imaging or cytobiochemical findings) consistent with infection. As the EORTC/IFICG and NIAID/MSG definitions do not provide specific indications for the clinical interpretation of Trichosporon sp. recovery from respiratory tract specimens, we have adapted the criteria for diagnosis of “probable” pulmonary infection proposed by Girmenia and collaborators (62). These criteria require the following evidence for diagnosis of probable pneumonia: the presence of pulmonary infiltrates and recovery of Trichosporon species from sputum or bronchoalveolar lavage (BAL) fluid samples in the absence of other pathogens causing opportunistic infections. Microbiological diagnosis of superficial and invasive trichosporonosis classically relies on culture findings as well as the identification of fungal elements compatible with Trichosporon spp. (hyphae, pseudohyphae, arthroconidia, and blastoconidia) in wet mount and/or tissue biopsy specimens. Although demonstration of fungal organisms in tissue and cultures is considered the “gold standard” for defining invasive infections, obtaining biopsy specimens may be difficult for some patients. Consequently, nonculture methods for the diagnosis of invasive mycosis are increasingly needed to surpass the limitations of test sensitivity, delayed results, and problems in distinguishing between colonized and truly infected patients. Antigen Detection The detection of (133)-␤-D-glucan (BDG) has been used extensively for early diagnosis of invasive fungal infections, including candidiasis and aspergillosis (136). There are no reports in the English language literature specifically addressing the sensitivity and specificity of BDG measurement in the diagnosis of invasive trichosporonosis. However, a recent publication reported epidemiological and clinical aspects related to 33 cases of Trichosporon fungemia in Japan (179). In that study, only 50% of patients had a single test positive for BDG at the time of admission, and few had an antigen-positive test

689

prior to their positive blood cultures. Therefore, based on the experience of this series of cases, it appears that the measurement of BDG levels may have serious limitations for the early diagnosis of invasive trichosporonosis. It is well known that Trichosporon strains have GXM in their cell wall, which can lead to a cross-reaction with C. neoformans antigens in sera of patients with invasive trichosporonosis (120). According to Lyman and collaborators, all Trichosporon strains can produce measurable levels of GXM antigen, and thus the detection of anticryptococcal cross-reactive antigen may be an useful tool in the early diagnosis of Trichosporon infections (109). However, unlike for C. neoformans, the structural and serological properties of Trichosporon GXM have been poorly investigated (56). Therefore, considering the limited clinical information available on this topic, further clinical studies are necessary to confirm if the detection of Trichosporon GXM by using the Cryptococcus antigen test has potential for screening patients with invasive trichosporonosis.

PCR-Based Methods More recently, PCR-based methods and flow cytometry assays have been developed to diagnose invasive infection due to Trichosporon species. Though these new assays are not yet standardized to be used routinely in clinical settings, they represent important strategies for future clinical validation. Nakajima et al. used a PCR technique to amplify the fungal ITS region. Total DNA was extracted from a lung biopsy specimen taken from a patient suffering from lung cancer. The ITS amplicon was sequenced, and this fragment identified T. asahii as the causative agent of the infection (131). Fungal DNA can be extracted and detected not only from fresh tissues but also from paraffin-embedded tissue sections. Sano et al. (158) extracted DNAs from 30 different tissue sections obtained from major organs of 3 autopsy cases of disseminated trichosporonosis. The fungemic patients had been previously diagnosed by blood cultures and analysis with the Vitek 2 compact system to be infected by T. asahii. Pan-species and species-specific primer sets were used in nested PCRs to amplify different amplicon sizes of the rDNA region: 170 bp, 259 bp, and 412 bp. Out of 30 tissue samples tested, 20 were positive for Trichosporon sp. infection by hematoxylin and eosin (H&E) staining, and 22 were positive by detection using Grocott’s stain. PCR amplification of the 170-bp fragment was positive in 20 of 22 samples, whereas only 12 of the 22 were positive for the 259-bp fragment. All of the 30 samples were PCR negative for the 412-bp fragment. Of the 8 samples that were negative by Grocott staining, only 1 sample was positive for both the 170- and 259-bp bands. The authors also verified that the efficiency of fungal detection by PCR decreased with longer periods of tissue fixation. After 1 day of fixation, only one-third of the collected samples were positive by both Grocott staining and amplification of the 170- and 259-bp bands. Longer tissue fixation of between 6 and 21 days considerably diminished the amplification of the 259-bp fragment (2 of 12) (158). Trichosporon spp. can also be detected from biological fluids by using PCR assays. Nagai et al. developed a nested PCR assay to amplify part of the T. asahii 28S rDNA from a collection of 11 serum samples from patients with histologically diagnosed disseminated trichosporonosis. Of the 11 samples

690

COLOMBO ET AL.

CLIN. MICROBIOL. REV.

FIG. 1. Schematic structure of the rRNA gene of Trichosporon spp. ITS, internal transcribed spacer; IGS, intergenic spacer region. The total length of the rDNA locus is approximately 7,850 bp, and the IGS1 region comprises 485 bp.

tested, 7 (64%) were PCR positive and 6 were positive for the GXM antigen (129). Later, Sugita and collaborators used species-specific primers to amplify the rDNA region in nested PCRs for T. asahii diagnosis. A total of 11 serum samples from 7 different patients with deep-seated trichosporonosis confirmed after autopsy were tested by PCR assay and latex agglutination (LA) test for detection of Trichosporon spp. T. asahii DNA was detected by PCR from 9 of 11 samples, with only 7 samples also positive in the LA test (173). Hosoki and collaborators (79) were able to detect breakthrough Trichosporon infection using serum PCR with a patient suffering from cytophagic histiocytic panniculitis and neutropenia. Those authors reported a study in which the patient’s serum was screened every other week to PCR amplify bands related to fungal DNA. It was observed that until the third week after a cord blood transplant, the patient had no fungemia. On day 28, when the patient had been receiving prophylactic voriconazole for 17 days, T. asahii DNA was detected in blood samples, even though blood cultures remained negative. At that time, the patient had no symptoms, and BDG serum titers were normal. T. asahii DNA was detected by PCR in this patient’s blood until day 39; the patient was then cured of the infection. To improve the detection and identification of fungal infections in biological samples, high-throughput technologies, such as PCR with pan-species primer sets and DNA microarray, have been applied. Microarray assays were combined with multiplex PCR of unique ITS1, 18S, and 5.8S sequences and consecutive DNA microarray hybridization to diagnose 14 major fungal pathogens, including T. asahii. Ninety-one samples of blood, bronchoalveolar lavage fluid, and tissues were analyzed from 46 neutropenic patients representing 5 cases with no fungal infection and 5 cases with proven, 3 cases with probable, and 33 cases with possible IFI. In the single case of invasive trichosporonosis included in this series, T. asahii could be positively detected in blood and BAL fluid samples by the microarray assay (166). Bottles of blood cultures collected from patients with invasive trichosporonosis have also been used to validate PCR assays as a tool for the diagnosis of fungemia and the further identification of the causative fungal species. Hsiue et al. developed an oligonucleotide array system targeting the ITS1 or ITS2 region to analyze 116 fungus-positive blood cultures. This system could identify 16 genera among 77 yeast species and was able to detect and identify 2 samples of T. asahii in blood cultures, compared to only one positive identification of Trichosporon spp. by phenotypic methods (82). More recently, Nagano and collaborators compared culturebased methods with molecular techniques for the detection of fungi in 77 adult cystic fibrosis patients. Sputum samples were collected for DNA extraction, and the ITS region was ampli-

fied for sequencing analysis. Trichosporon spp. were identified in 2 of 77 samples by molecular techniques, whereas only one of those two samples was identified by culture methods (130). Luminex xMAP Technology A novel flow cytometric technique called Luminex xMAP has been developed for the detection of medically important species of fungi. This assay is based on the use of PCR-biotinylated amplicon target DNA that is inoculated into a microsphere-bead mixture containing species-specific probes of interest. By adding a reporter molecule (streptavidin R-phycoerythrin), hybridized species-specific amplicons captured by complementary nucleotide sequences on microsphere beads can be recognized by the fluorescence of the reporter molecules. Landlinger and collaborators (103) have used the Luminex xMAP technology to identify fungal species in samples from peripheral blood, blood cultures, pulmonary infiltrate biopsy specimens, bronchoalveolar lavage fluid, and bronchotracheal secretions and from isolated fungal strains. This system utilized 2 pan-Trichosporon probes designed to hybridize to the ITS2 region, which is highly variable among the genomes of individual fungal species, and it detected at least four clinically relevant Trichosporon species (T. asahii, T. inkin, T. cutaneum, and T. beigelii). Those authors reported that from one sample of paraffin-embedded tissue, a Trichosporon sp. was detected by specific hybridization signals, and following further sequencing analysis, this species was identified as T. cutaneum. Not only is this technology useful for analysis of biological samples, it also has potential for the identification of Trichosporon spp. from culture isolates by using probes that recognize not only the ITS region and intergenic spacer 1 (IGS1) but also the D1/D2 region of the 28S rDNA (Fig. 1). Diaz et al. (41) used the Luminex 100 xMAP assay to identify 39 strains of different Trichosporon species. They showed that the ITS and D1/D2 regions were sufficient to discriminate between species and that the IGS region could differentiate closely related species such as T. asahii, T. japonicum, and T. asteroides. New Perspectives on Microscopic Imaging Diagnosis Microscopic imaging analysis has also been improved in order to increase the ability to prove deep-seated Trichosporon spp. infections. Obana et al. (135) compared the performances of different histopathological stains for identifying fungal cells in tissues from 9 autopsy cases, including 3 cases of disseminated trichosporonosis, 3 cases of gastric candidiasis, 2 cases of pulmonary aspergillosis, and 1 case of pulmonary cryptococcosis. Those authors reported that Trichosporon cells (T. asahii) were weakly detectable within tissue sections compared with the ability to detect other fungal species when stained with conventional Grocott’s stain. Using a mucicarmine stain, Tri-

VOL. 24, 2011

chosporon could be weakly stained within the tissues, whereas Candida and Aspergillus were negatively stained. A colloidal iron stain, which reacts with Cryptococcus capsules, showed clear staining of Trichosporon cell walls as well as Cryptococcus capsules. A periodic acid-methenamine-silver stain (PAM) appeared to best discriminate fungal elements of Trichosporon spp. in tissue sections. Those authors used transmission electron microscopy to observe sections stained with Grocott’s stain and diluted PAM to evaluate putative differences in cell wall silver deposition between Candida and Trichosporon strains. This strategy was useful for identifying differences in hyphal size as well as laminar deposition of silver granules in the cell wall. In conclusion, the authors suggested that histological staining procedures and electron microscopy may have potential as a tool for discriminating tissue infections by Candida and Trichosporon species. Phenotypic Identification of Trichosporon Species The taxonomy of Trichosporon spp. has been completely rewritten, and the genus now includes 50 species (139), at least 16 of which have clinical relevance. This high number of new species presents a large variability in terms of virulence, clinical manifestations in humans, and antifungal susceptibility. Consequently, before defining strategies to treat and prevent superficial and fungal infections caused by Trichosporon spp., it is necessary to accurately identify the causative microorganism at the species level. Several methods to identify Trichosporon species have been reported, including morphological and biochemical tests as well as molecular tools. Despite the fact that phenotypic methods are more suitable for routine use in general microbiology laboratories, the accuracy of the identification of Trichosporon spp. seems to be limited. Molecular methods are more precise in species identification but are costly for routine laboratory use (151, 172). It is important to note that both Trichosporon and Geotrichum species are able to produce arthroconidia. In clinical laboratory analysis, when arthroconidia are visualized the urease test is recommended. Unlike Geotrichum spp., all species of the Trichosporon genus are able to hydrolyze urea (39). Although these genera are phenotypically similar, they are genetically very distinct. The ITS nucleotide sequences are less than 80% similar (33). Subsequently, specific diagnostic PCR can also be performed to differentiate these two genera (175). Phenotypic methods for Trichosporon species identification are based on the characterization of micromorphological aspects of colonies as well as biochemical profiling. Performing a slide microculture to search for arthroconidia is a very useful tool for the screening of Trichosporon spp. However, other morphological aspects and biochemical tests do not allow the complete identification of Trichosporon isolates at the species level (151, 172). Indeed, by comparing the experiences of 3 different laboratories in evaluating the biochemical and physiological characteristics of 6 clinically relevant Trichosporon spp., significant discrepancies in the results generated by testing the same organisms were observed in different studies (24, 39, 144, 175). Therefore, one can conclude that classical methods for yeast identification have limited accuracy and reproducibility for Trichosporon spp. identification.

TRICHOSPORON SPP. AND TRICHOSPORONOSIS

691

Despite limitations, several commercial nonautomated and automated systems have been used in the identification of Trichosporon spp. Notwithstanding the problems for biochemical tests mentioned above, it is also important to emphasize that many of these methods do not include new taxonomic categories in their databases. Consequently, the identification of the genus Trichosporon is oversimplified due to incomplete databases and classification keys. The nonautomated commercial methods most commonly used for yeast identification in clinical laboratories are summarized below. 1. API 20C AUX (bioMe´rieux, Mercy l’Etoile, France). The API 20C AUX method is based on the evaluation of the ability of the fungus to assimilate 19 carbon sources. It requires additional tests, such as macro- and micromorphological observation of colonies and a urease production test. The results are read at 48 or 72 h. The current database includes only 3 species: T. asahii, T. inkin, and T. mucoides (48, 72, 144, 146). 2. ID 32C (bioMe´rieux, Mercy l’Etoile, France). The ID 32C method is based on the evaluation of the ability of the fungus to assimilate 24 carbon sources and 5 organic acids, evaluation of yeast sensitivity to cycloheximide, and esculin tests. It requires additional observation of the macro- and micromorphology of the colonies. This method shows inconsistent identification, and the database includes only T. asahii, T. inkin, and T. mucoides (18, 144, 146). 3. RapID Yeast Plus system (Innovative Diagnostic System, Norcross, GA). The RapID Yeast Plus system is based on the observation of 13 enzymatic hydrolysis substrates and assimilation of 5 carbon sources for the identification of clinically important yeasts (48). Kitch et al. (97) analyzed the ability of this technique to identify 304 clinical yeast isolates, 3 of which were Trichosporon spp., within 4 h. Two out of 3 isolates were identified exclusively as Trichosporon beigelii, though 1 isolate was misidentified as C. neoformans. The automated systems for yeast identification most commonly used in clinical laboratories are summarized below. In general, all of the automated systems have severe limitations regarding the number of Trichosporon species listed in their databases. 1. Vitek Systems (bioMe´rieux, Vitek, Hazelwood, MO). The Vitek Systems system is based on nitrogen and carbon assimilation tests, enzymatic tests, and evaluation of yeast sensitivity to cycloheximide. The test readings are performed spectrophotometrically by the automated system after 24 and 48 h of incubation. The current database (Vitek 2) lists only T. asahii, T. inkin, and T. mucoides. Not only is this system limited by the number of species included in the database, but inconsistent results have been reported for the identification of T. inkin and T. asteroides, being erroneously identified as T. asahii by this system (2, 19, 43, 52, 83). 2. Baxter Microscan (Baxter Microscan, West Sacramento, CA). The Baxter Microscan method can generate final reports of yeast identification in 4 h by evaluating the

692

COLOMBO ET AL.

enzymatic profile of each organism tested. Fluorometric and spectrophotometric readings are taken and enriched by data generated by micro- and macromorphological observation of colonies. This system is considered to be less accurate than the Vitek system described above, and several authors have experienced difficulties in the correct identification of Trichosporon spp. using this method due to limitations in its database. To our knowledge, there has not been an update to the Microscan database since the original publications; therefore, the database still refers to all Trichosporon spp. as T. beigelii (102, 168). Most papers addressing the accuracy of commercial yeast identification methods provide very little data on specific Trichosporon species. Moylett and collaborators claimed to identify 2 isolates of Trichosporon pullulans as the cause of chronic granulomatous disease by using the Vitek Yeast Biochemical card (128). At that time, this card version had only T. beigelii and T. pullulans in its database. To confirm their findings, Holland et al. examined the same two strains by amplification and sequencing of their ITS regions (78). BLAST analysis revealed that both yeasts initially identified phenotypically as Trichosporon pullulans were genotypically identified as Cryptococcus adeliensis. Ahmad and collaborators used the Vitek 2 system to identify a collection of clinical strains and found 29 isolates that they first identified as T. asahii. After all 29 identifications were checked by sequencing the ITS regions, 4 strains were reassigned as T. asteroides (2). More recently, Leaw et al. (105) analyzed the accuracy of phenotypic tests and molecular markers (ITS1, ITS2, and 28S D1/D2 regions) for the identification of 373 medically relevant yeasts encompassing 86 species. A total of 74 strains were simultaneously identified by the API ID32C system and molecular methods. Of note, 27 Trichosporon strains representative of 9 different species (T. aquatile, T. asahii, T. cutaneum, T. debeurmannianum, T. dermatis, T. inkin, T. jirovecii, T. mucoides, and T. pullulans) were included in the analysis. The first finding of this study was that examination of the ITS2 region allowed for accurate identification of the yeast species, while the D1/D2 sequences were too conserved and could not discriminate between T. dermatis and T. mucoides. In addition, the authors reported that 4 out 6 T. dermatis clinical isolates were misclassified by biochemical methods as T. cutaneum. The limited ability of biochemical tests to provide reliable identification of clinical isolates of Trichosporon spp. was recently documented by Rodriguez-Tudela et al. (151). They compared the identification of 49 isolates of Trichosporon spp. using the amplification and sequencing of the ITS and intergenic spacer (IGS) regions with that of 4 different biochemical tests, including fermentation of different carbon sources, growth on nitrogen sources, growth at various temperatures, and the ability to hydrolyze urea. Not surprisingly, 26 isolates (53%) were misclassified at the species level according to sequencing analysis. Out of 15 T. asahii isolates, only 8 were correctly identified by biochemical tests, while 6 were identified as T. mucoides and 1 as T. ovoides. All isolates belonging to the T. jirovecii, T. japonicum, T. montevideense, and T. domesticum species were not able to be identified by classical

CLIN. MICROBIOL. REV.

identification methodology. However, 6 T. inkin isolates out of 7 were correctly identified by biochemical tests. Commercial systems based on phenotypic tests to provide identification of Trichosporon spp. are very limited in their accuracy because of either the limited number of species included in their databases or the inconsistent ability of their biochemical tests to accurately identify all 50 potential species within the genera. Considering the limitations of the phenotypic methods used to identify Trichosporon spp. at the species level, it is easy to understand why most reports refer only to the genus Trichosporon without determining the species or simply identify clinical isolates as T. asahii or non-T. asahii. The lack of accurate laboratory tools for the complete identification of Trichosporon strains in clinical laboratories impairs the understanding of unique epidemiological and clinical features of these strains. This technical deficiency also impairs the ability to understand differences in clinical responses to conventional antifungal therapy, possibly related to the medically important species of this genus, previously named T. beigelii (10, 141, 185, 186). Molecular Targets and Identification of Trichosporon Species in Cultures PCR-based methods. DNA-based methods have been extensively used for the accurate identification of Trichosporon spp. (142). Indeed, the evaluation of specific nucleotide sequences can be a precise method to resolve taxonomic problems generated by the inconsistent phenotypic identification of Trichosporon species. In this regard, ribosomal genes represent particularly consistent evaluative markers and include alternating conserved regions (D1/D2 region of the 28S rDNA) and variable regions (ITS and IGS1 regions) that may be useful for species identification and phylogenetic studies (Fig. 1) (171, 175). Apparently, a combination of at least 2 of those markers should be used for the proper molecular identification of Trichosporon strains and analysis of their phylogenetic relationships. Sugita et al. (175) constructed a phylogenetic tree using the small-subunit (SSU) region sequences of rDNAs from different pathogenic yeasts obtained from DNA libraries. The primer pair TRF and TRR, which amplifies part of the SSU region, was designed for the specific identification of the genus Trichosporon because these oligonucleotides do not amplify conserved regions in the ribosomal genes of other medically important yeasts besides Trichosporon. Subsequently, Sugita et al. (174) have sequenced and analyzed the interspacer region (ITS1 and ITS2) genes of the rDNA from Trichosporon spp. and proposed 17 species and five varieties for this genus. Therefore, these authors concluded that the six medically relevant species could be accurately identified by their ITS sequences. However, Sugita et al. (172) also analyzed the sequences of IGS1, which is localized between the 26S and 5S rDNA genes, in 25 isolates of Trichosporon. The IGS1 regions ranged in size from 195 bp to 704 bp. Comparative analysis of the nucleotide sequences suggested higher variation in the IGS1 region than in the ITS region. Therefore, the use of ITS region sequencing was considered unsuitable for the identification of the large number of recently described species of Trichosporon. In ad-

VOL. 24, 2011

dition, these authors were also able to identify 5 different genotypes of T. asahii among 43 strains (172). Following this work, several others have reported the use of the IGS region to accurately identify Trichosporon spp. in cultures from several biological samples, such as skin, nails, urine, sputum, biopsy samples, and blood (25, 89, 104, 151, 169). Based on the ITS1 and ITS2 sequences, Leaw et al. (104) developed an oligonucleotide array to identify 77 species of clinically relevant yeasts belonging to 16 genera. Probes for each yeast species were designed and included probes to identify T. aquatile, T. asahii, T. cutaneum, T. inkin, and T. pullulans, as well as a pan-probe for the identification of T. asahii, T. aquatile, and T. inkin. Those authors screened a collection of 452 yeast strains and verified that the array had 100% sensitivity and 97% specificity. There were 17 Trichosporon sp. type strains and 4 T. cutaneum clinical isolates included in the collection to validate the array and 4 other species, (T. dermatis, T. jirovecii, T. mucoides, and T. ovoides) to test the assay specificity. The authors concluded that all Trichosporon spp. tested were correctly identified except for T. ovoides, which cross-hybridized with the pan-probe (104). More recently, Makino et al. (112) used a real-time quantitative PCR (qPCR) assay to identify and quantify 9 yeast species, including T. asahii and T. jirovecii, in dairy product samples. They amplified conserved sequences of 28S rDNA D1/D2 domains with 6 primer sets and were able to identify all isolates that artificially contaminated fermented milk within a 5-h assay. These tools and DNA markers were useful for yeast identification and could possibly be used as a first screening tool for Trichosporon species in biological samples. Other molecular markers have been tested for use in the identification of Trichosporon species. Biswas et al. (20) analyzed the sequences of the mitochondrial cytochrome b (cyt b) genes from 23 different fungal strains representing 42 species. They amplified a 396-bp fragment and observed that there were 141 variable nucleotide sites (35.6%) among Trichosporon strains. Twenty-two strains (from 11 different species) contained introns in their sequences. Analysis of the deduced protein sequences of the 396-bp fragments showed 34 variable amino acid sites (25.75%). T. domesticum, T. montevideense, T. asahii, T. asteroides, T. gracile, and T. guehoae all had identical amino acid sequences. Phylogenetic analysis revealed that all species of Trichosporon except T. montevideense and T. domesticum included species-specific cyt b genes. Using this approach, two isolates were then identified: Trichosporon sp. strain CBS 5581 was identified as T. pullulans, and the clinical isolate IFM 48794 was identified as T. faecale (20). Proteomics as a tool for fungal identification. During the past few years, protein fingerprints have been used as genus and species signatures to discriminate among isolates and identify pathogens (58). Protein fingerprint analysis by mass spectrometry (MS), in which the protein content of a certain sample within a complex mixture is analyzed, is a widely used technique in proteomics studies. Mass spectrum profiles are acquired by mixing the samples with a matrix in which peptides are ionized using a laser. Thereafter, peptides are expelled from the metal target by high voltage and travel in a vacuum tube to be finally analyzed according to their mass/charge ratio (m/z). This technique typically generates spectra of the most abundant proteins, ranging from

TRICHOSPORON SPP. AND TRICHOSPORONOSIS

693

1,000 to 20,000 Da. The process to identify microorganisms in culture, including automated identification, is based on the matching of peaks from fingerprints deposited in a database, e.g., the matrix-assisted laser desorption ionization– time of flight (MALDI-TOF) MS BioTyper system (Bruker Daltonics), with the peaks of the spectrum generated from the clinical sample. The more similar two spectra are, the more reliable is the match, and therefore, genus or species identification is obtained (21). The MALDI BioTyper 2.0 software possesses more than 3,700 mass spectrum profiles in its database. Of those profiles, 274 are from fungi and 11 are related to Trichosporon spp. Of these 11, 6 are T. asahii, T. cutaneum, T. debeurmannianum, T. inkin, T. mucoides, and T. ovoides and 2 are Trichosporon sp. Another feature of this method is the time required for sample processing. One isolate can be processed and identified in less than 2 min, making this tool the next generation in microorganism identification (58). Marklein and collaborators (115) have identified 285 isolates of medically important yeasts, including 3 isolates of T. cutaneum, using MALDI-TOF MS. Overall, the system identified 247 of the clinical isolates (92.5%); the remaining 20 isolates were identified only after complementation of the database with protein spectra representative of all type strains. All 3 isolates of T. cutaneum were properly identified by the API ID 32C system but not by the MALDI-TOF MS due to the absence of a reference protein spectrum characteristic for this species. After generation of a reference type spectrum for T. cutaneum, all 3 isolates were appropriately identified. Recently, Bader et al. (13) tested the accuracy of MALDI-TOF MS using 2 different commercially available systems, MALDI BioTyper2 (Bruker Daltonics) and Saramis (AnagnosTec), for the identification of 1,192 clinical yeast isolates previously identified by morphological characterization and biochemical tests (API 20 C AUX and ID 32 C galleries). The congruence among all three identification methods was 95.1%, and all 4 isolates of T. asahii were properly identified by these three procedures. Stevenson et al. (167) constructed a yeast mass spectrum library using the MALDI BioTyper2 (Bruker Daltonics) software. One hundred twenty-four type strains were initially evaluated for construction of the protein spectrum database library, but 15 were removed due to unacceptable spectra or discordant identification results after DNA sequencing, which occurred with 1 T. asahii strain and 3 T. ovoides strains. In order to validate the accuracy of the database library, 194 clinical isolates (23 species covering 6 genera) were tested, and 192 (99.0%) of the clinical isolates were properly identified by MALDI-TOF MS (ID score of ⬍1.8), including 4 isolates of T. asahii, 1 of T. coremiiforme, and 3 of T. mucoides. MALDITOF MS should be considered a reliable technique to rapidly identify fungal species and may be used as an alternative to PCR-based methods, with the additional advantage of the lack of complicated DNA extraction (167). It is therefore evident that the use of molecular methods to evaluate specific DNA sequences or proteins fingerprints is now considered mandatory to accurately identify strains belonging to the genus Trichosporon. Small clinical laboratories unable to run those tests should send their strains to reference

694

COLOMBO ET AL.

CLIN. MICROBIOL. REV.

FIG. 2. Distribution of T. asahii genotypes, based on IGS1 sequences of clinical isolates, in the United States, Spain, Brazil, Thailand, Japan, and Turkey (25, 89, 119, 152, 169, 172).

laboratories to have the Trichosporon strains properly identified at the species level, particularly when such strains are obtained from deep-seated infections. Genotyping of Trichosporon spp., with an emphasis on the geographic distribution of T. asahii genotypes. T. asahii is the most commonly isolated species from patients with invasive trichosporonosis and presents up to 9 different genotypes described around the globe, using the IGS1 sequence as the molecular marker (24, 119). Sugita et al. were the first to describe 5 T. asahii genotypes, based on the diversity of the IGS1 sequences exhibited by T. asahii strains obtained from Japan, the United States, and Brazil (172). They found that among isolates obtained in Japan, 26 (87%) were genotype 1 or 3, while the others belonged to genotypes 2 and 4. The 13 T. asahii strains originally from the United States were representative of either genotype 3 or genotype 5, and the single Brazilian isolate was identified as genotype 3. No genotype 1 strains were found among the 13 North American strains tested. Unlike the observations made with patients with superficial and invasive trichosporonosis, Sugita et al. reported that most T. asahii strains isolated from the homes of patients suffering with summer-type hypersensitivity pneumonitis were genotype 3, instead of genotype 1, which is predominant in patients. In the same study, those authors described 2 new genotypes of T. asahii, genotypes 6 and 7, both documented in patients with fungemia (169). Rodriguez-Tudela et al. (152) also evaluated sequence polymorphisms of the IGS1 regions of T. asahii strains isolated from patients in Argentina, Brazil, and Spain. The authors were able to recognize 6 different genotypes within the collection of 18 T. asahii strains. The majority of the strains were representative of genotypes 1 and 5. Spanish strains exhibited all other genotypes, with the exception of genotype 2, whereas the 5 South American isolates belonged to genotypes 1 and 6 (152). Chagas-Neto and collaborators analyzed the genotype distribution of 15 T. asahii isolates obtained from blood samples from patients in Brazil and found that the majority of these strains belonged to genotype 1 (86.7%). Only one isolate each belonged to genotypes 3 and 4, representing the first description of genotype 4 reported in a patient in South America (25). Recently, two papers described the geographic distribution pattern of T. asahii genotypes in Turkey and Thailand based on

IGS1 sequences (Fig. 2). Kalkanci et al. reported that among 87 T asahii clinical isolates from Turkey, genotype 1 was represented by 69 strains (79.3%), followed by genotypes 5 (7 strains, 8.0%), 3 (6 strains, 6.9%), 6 (3 strains, 3.4%), and 4 (1 strains, 1.1%) (89). By examining the genotyping of T. asahii strains performed worldwide, those authors noted that genotype 7 has been found only in Japan and that genotype 8 and the novel genotype 9 were described only in Turkey. Mekha et al. evaluated 101 T asahii strains in Thailand and found that genotype 1 (45 strains, 44.5%) was predominant, followed by genotypes 3 (35 strains, 34.7%) and 6 (18 strains, 17.8%) (119). Figure 2 summarizes the worldwide distribution of T asahii strains obtained only from patients with superficial and invasive Trichosporon infections (25, 89, 119, 152, 169, 172). It is important to note that, with the exception of Turkey, Thailand, and Japan, a limited number (usually ⬍30) of T. asahii strains have been genotyped in most studies published in the English language literature. Genotype 1 is the most abundant type of T. asahii strain in all regions, ranging from approximately 45 to 80%, except for the United States where it has never been identified. Genotypes 3 and 5 predominate in U.S. medical centers. Further studies are needed to clarify whether different genotypes of T. asahii isolates may present different features in terms of virulence, tissue tropism, or antifungal susceptibility. ANTIFUNGAL SUSCEPTIBILITY TESTING FOR TRICHOSPORON SPP. AND PRINCIPLES OF THERAPY Despite the increasing occurrence of invasive Trichosporon infections refractory to conventional antifungal drugs, there are few studies investigating the in vitro susceptibilities of clinical strains of Trichosporon spp. to antifungal compounds. Difficulties in species identification within the genus and the lack of standardized sensitivity tests in vitro contribute to the limited information available on this topic (24, 140). Antifungal Susceptibility Tests The Clinical and Laboratory Standards Institute (CLSI) document describing antifungal susceptibility testing of yeasts does not specifically address the genus Trichosporon. However, most available studies evaluating the susceptibility of Trichosporon

VOL. 24, 2011

TRICHOSPORON SPP. AND TRICHOSPORONOSIS

spp. to antifungal drugs in vitro use the CLSI broth microdilution method currently (2008) standardized for Candida spp. and C. neoformans (10, 132) or an adaptation of the European Committee for Antimicrobial Susceptibility Testing (EUCAST) broth microdilution method, a recommendation originally proposed for the genus Candida (36). Despite the fact that the CLSI methodology may be successfully adapted to test Trichosporon strains, several authors have suggested that the broth microdilution method is not satisfactory for detecting isolates resistant to amphotericin B, as it generates a narrow MIC variation within different clinical strains. Therefore, MIC breakpoints have not yet been established for amphotericin B assays (41, 49). The majority of the studies examining sensitivity tests of Trichosporon spp. use the outdated nomenclature T. beigelii (28, 75, 141, 185). Therefore, sensitivities to antifungal drugs within different species of the genus Trichosporon are largely still not investigated. After the resolution of the genus Trichosporon, some authors have suggested that T. asahii is more resistant to amphotericin B and more sensitive to triazoles than other Trichosporon species (80, 89, 190). Rodriguez-Tudela et al. (151) accurately identified 49 Trichosporon clinical isolates using IGS region sequencing and tested their susceptibility to antifungal drugs. Those authors demonstrated that all T. asahii isolates tested had amphotericin B MICs of ⱖ2 ␮g/ml. They also observed that the majority of T. coremiiforme and T. faecale strains were also resistant to amphotericin B, whereas the other species tested had MICs of ⬍1 ␮g/ml. In a recent study coordinated by the same group, voriconazole and posaconazole were shown to be the 2 new antifungal drugs with the best in vitro antifungal activity against most Trichosporon spp. (8). Chagas-Neto et al. (25) evaluated the in vitro activities for 22 bloodstream Trichosporon strains using the CLSI broth microdilution test. The authors found that the geometric means of MICs generated by triazoles against Trichosporon spp. were generally lower than those values obtained with amphotericin B. Voriconazole generated the lowest MIC values against all clinical strains (25). Ruan et al. (154) evaluated the in vitro activities of 9 different antifungal drugs against 43 clinical isolates of Trichosporon species using broth microdilution. Voriconazole exhibited excellent in vitro activity against most of the strains tested, including isolates resistant to fluconazole. All isolates appeared to be resistant in vitro to all echinocandins. The authors concluded that most strains exhibited relatively high MIC values against amphotericin B. Antifungal Therapy Despite the increasing relevance of the genus Trichosporon in contemporary medicine, the treatment of patients with trichosporonosis remains a challenge, as there are few data available on the in vitro and in vivo activities of antifungal drugs against clinically relevant species of the genus. Superficial infections occasionally respond to topical antifungal agents, but white piedra may reoccur after suspension of the antimycotic. Consequently, some studies suggest that treatment of this disease should consist of hair removal followed by topical and/or oral antifungal therapy (96).

695

There are increasing data suggesting that amphotericin B has limited in vitro and in vivo activity against Trichosporon spp., including T. asahii strains (183, 193). Amphotericin B MIC values against Trichosporon spp., particularly T. asahii, are generally incompatible with serum levels normally achievable in patients receiving a conventional formulation of this polyene. Although some isolates can be inhibited by low concentrations of amphotericin B, fungicidal activity has not been observed in animal models or in neutropenic patients (3, 24, 25, 143, 179, 186, 189, 190). In a study of 25 neutropenic patients who developed systemic trichosporonosis and were treated with amphotericin B, only four survived (81). Girmenia et al. reported the clinical outcomes for 55 patients with hematological diseases and disseminated trichosporonosis who were treated with amphotericin B. A clinical response to amphotericin B was documented in only 13/55 (24%) of the patients evaluated (62). Triazoles seem to have better in vitro and in vivo antifungal activities against Trichosporon spp. than amphotericin B. Fluconazole was used as the initial therapy in 85% of 19 patients with invasive trichosporonosis documented in a single medical center in Taiwan, and the mortality rate was 42% (154). Suzuki et al. recently evaluated the clinical and therapeutic aspects of 33 cases of Trichosporon fungemia documented in patients with hematological malignancies. The authors clearly demonstrated that survival was longer for patients treated with an azole than for those treated with other drugs (179). Voriconazole also has excellent in vitro activity against Trichosporon strains and may be useful for treating patients with trichosporonosis, including cases of acute leukemia and myelodysplasic syndrome with disseminated infection (12, 57, 106, 118, 140, 160). Regarding the therapeutic role of 5-fluorocytosine (5FC) in trichosporonosis, the available data are limited and controversial. In vitro data suggest that a large proportion of Trichosporon strains may be less susceptible or resistant to 5FC (25, 119). In contrast, some authors report good results in the treatment of trichosporonosis with combination therapy with 5FC and amphotericin B (62). Consequently, at this time, we do not have substantial clinical data to safely recommend the utilization of 5FC to treat invasive trichosporonosis. Finally, echinocandins alone have little to no activity against Trichosporon spp. and are not recommended for trichosporonosis treatment (188). Breakthrough Trichosporon infections have been reported in patients treated with echinocandins (caspofungin and micafungin) (35, 47, 188). Interestingly, a combination of echinocandin with amphotericin B or azoles appears to have some in vitro and in vivo synergistic antifungal effects (14, 161). SUMMARY AND CONCLUSION Trichosporon spp. are basidiomycetous yeast-like anamorphic organisms, found in tropical and temperate areas of the globe and are widely distributed in nature. Although most Trichosporon strains isolated in clinical laboratories are related to episodes of colonization or superficial infections, this fungus has been recognized as an opportunistic agent causing emergent, invasive infections in tertiary care hospitals worldwide.

696

COLOMBO ET AL.

Invasive Trichosporon spp. have been documented mostly in cancer patients and in critically ill patients exposed to multiple invasive medical procedures. It is possible that the ability of Trichosporon strains to adhere to and form biofilms on implanted devices may account for the progress of invasive trichosporonosis, since this ability promotes escape from drugs and host immune responses. In addition, the presence of glucuronoxylomannan (GXM) in the cell walls of Trichosporon spp. and their ability to produce proteases and lipases are all factors likely related to the virulence of this genus. The outdated taxon T. beigelii was replaced by several separate species, and the taxonomy of the genus was progressively modified by powerful molecular tools able to discriminate between phylogenetically closely related species. Currently, 50 different species of Trichosporon have been described by different authors, and 16 can cause human disease. Globally, T. asahii is the most common species causing invasive trichosporonosis and presents 9 different genotypes, based on the IGS1 sequence, that exhibit substantial variability in their geographic distribution worldwide. Phenotypic methods for Trichosporon species identification are based on the characterization of micromorphological aspects of colonies and biochemical profiling. Performing a slide microculture to search for arthroconidia and testing for urease production are useful tools for the screening of Trichosporon spp. However, morphological aspects and biochemical tests do not allow the complete identification of Trichosporon isolates at the species level. Consequently, molecular methods, including sequencing of the IGS region, are necessary to obtain accurate results for the identification of Trichosporon strains. Diagnosis of invasive trichosporonosis may be a challenge, and several molecular methods have been developed in the last 2 decades, including PCR-based methods, Luminex xMAP technology (a novel flow cytometry technique with potential for the detection of medically important species of fungi), and proteomics. Despite the increasing relevance of the genus Trichosporon in contemporary medicine, treating patients with trichosporonosis remains a challenge given that we have little data available on the in vitro and in vivo activities of antifungal drugs against clinically relevant species of the genus. In conclusion, disseminated trichosporonosis has been increasingly reported worldwide and represents a challenge for both diagnosis and species identification. Prognosis is limited, and antifungal regimens containing triazoles appear to be the best therapeutic approach. In addition, removal of central venous lines and control of underlying conditions should be considered to optimize clinical outcomes.

ACKNOWLEDGMENTS This work was supported in part by the Fundac¸˜ao de Amparo a Pesquisa do Estado de Sa˜o Paulo (FAPESP), Brazil (grants 2005/ 02006-0, 2005/04442-1, and 2007/08575-1), and the Conselho Nacional de Desenvolvimento Científico e Tecnolo ´gico (CNPq), Brazil (grant 308011/2010-4). A.C.B.P. received a postdoctoral fellowship from the Coordenac¸˜ao de Aperfeic¸oamento de Pessoal de Nível Superior (CAPES), Brazil (PNPD 02640-09-0). We report no conflicts of interest. We alone are responsible for the content and writing of the paper.

CLIN. MICROBIOL. REV. REFERENCES 1. Abdala, E., R. I. Lopes, C. N. Chaves, E. M. Heins-Vaccari, and M. A. Shikanai-Yasuda. 2005. Trichosporon asahii fatal infection in a non-neutropenic patient after orthotopic liver transplantation. Transpl. Infect. Dis. 7:162–165. 2. Ahmad, S., M. Al-Mahmeed, and Z. U. Khan. 2005. Characterization of Trichosporon species isolated from clinical specimens in Kuwait. J. Med. Microbiol. 54:639–646. 3. Anaissie, E., et al. 1992. Azole therapy for trichosporonosis: clinical evaluation of eight patients, experimental therapy for murine infection, and review. Clin. Infect. Dis. 15:781–787. 4. Anderson, J. M., and D. R. Soll. 1987. Unique phenotype of opaque cells in the white-opaque transition of Candida albicans. J. Bacteriol. 169:5579–5588. 5. Ando, M., K. Arima, R. Yoneda, and M. Tamura. 1991. Japanese summer-type hypersensitivity pneumonitis. Geographic distribution, home environment, and clinical characteristics of 621 cases. Am. Rev. Respir. Dis. 144:765–769. 6. Ando, M., et al. 1990. Serotype-related antigen of Trichosporon cutaneum in the induction of summer-type hypersensitivity pneumonitis: correlation between serotype of inhalation challenge-positive antigen and that of the isolates from patients’ homes. J. Allergy Clin. Immunol. 85:36–44. 7. Arai, T., S. Yusa, K. Kirimura, and S. Usami. 1997. Cloning and sequencing of the cDNA encoding lipase I from Trichosporon fermentans WU-C12. FEMS Microbiol. Lett. 152:183–188. 8. Araujo Ribeiro, M., A. Alastruey-Izquierdo, A. Gomez-Lopez, J. L. Rodriguez-Tudela, and M. Cuenca-Estrella. 2008. Molecular identification and susceptibility testing of Trichosporon isolates from a Brazilian hospital. Rev. Iberoam. Micol. 25:221–225. 9. Archer-Dubon, C., et al. 2003. Superficial mycotic infections of the foot in a native pediatric population: a pathogenic role for Trichosporon cutaneum? Pediatr. Dermatol. 20:299–302. 10. Arikan, S., and G. Hascelik. 2002. Comparison of NCCLS microdilution method and Etest in antifungal susceptibility testing of clinical Trichosporon asahii isolates. Diagn. Microbiol. Infect. Dis. 43:107–111. 11. Arikawa, T., et al. 2010. Galectin-9 expands immunosuppressive macrophages to ameliorate T-cell-mediated lung inflammation. Eur. J. Immunol. 40:548–558. 12. Asada, N., et al. 2006. Successful treatment of breakthrough Trichosporon asahii fungemia with voriconazole in a patient with acute myeloid leukemia. Clin. Infect. Dis. 43:39–41. 13. Bader, O., et al. 2010. Improved clinical laboratory identification of human pathogenic yeasts by matrix-assisted laser desorption ionization time-offlight mass spectrometry. Clin. Microbiol. Infect. in press. 14. Bassetti, M., et al. 2004. Trichosporon asahii infection treated with caspofungin combined with liposomal amphotericin B. J. Antimicrob. Chemother. 54:575–577. 15. Bayramoglu, G., M. Sonmez, I. Tosun, K. Aydin, and F. Aydin. 2008. Breakthrough Trichosporon asahii fungemia in neutropenic patient with acute leukemia while receiving caspofungin. Infection 36:68–70. 16. Behrend, G. 1890. Ueber Trichomycosis nodosa (Juhel-Re´noy): Piedra (Osorio). Klin. Wschr. 27:464–467. 17. Benson, P. M., N. A. Lapins, and R. B. Odom. 1983. White piedra. Arch. Dermatol. 119:602–604. 18. BioMe´rieux. 2006. ID 32 C—identification system for yeasts. bioMe´rieux, Marcy l’Etoile, France. 19. BioMe´rieux. 2008. Identification VITEK 2 YST CARD. bioMe´rieux, Marcy l’Etoile, France. 20. Biswas, S. K., L. Wang, K. Yokoyama, and K. Nishimura. 2005. Molecular phylogenetics of the genus Trichosporon inferred from mitochondrial cytochrome B gene sequences. J. Clin. Microbiol. 43:5171–5178. 21. Bruker Daltonics. 2008. MALDI Biotyper user manual, version 2.0 SR1. Bruker Daltonik GmbH, Bremen, Germany. 22. Cafarchia, C., D. Romito, C. Coccioli, A. Camarda, and D. Otranto. 2008. Phospholipase activity of yeasts from wild birds and possible implications for human disease. Med. Mycol. 46:429–434. 23. Cawley, M. J., et al. 2000. Trichosporon beigelii infection: experience in a regional burn center. Burns 26:483–486. 24. Chagas-Neto, T. C., G. M. Chaves, and A. L. Colombo. 2008. Update on the genus Trichosporon. Mycopathologia 166:121–132. 25. Chagas-Neto, T. C., G. M. Chaves, A. S. Melo, and A. L. Colombo. 2009. Bloodstream infections due to Trichosporon spp.: species distribution, Trichosporon asahii genotypes determined on the basis of ribosomal DNA intergenic spacer 1 sequencing, and antifungal susceptibility testing. J. Clin. Microbiol. 47:1074–1081. 26. Chakrabarti, A., et al. 2002. Generalized lymphadenopathy caused by Trichosporon asahii in a patient with Job’s syndrome. Med. Mycol. 40:83–86. 27. Chan, R. M., P. Lee, and J. Wroblewski. 2000. Deep-seated trichosporonosis in an immunocompetent patient: a case report of uterine trichosporonosis. Clin. Infect. Dis. 31:621. 28. Chaturvedi, V., R. Ramani, and J. H. Rex. 2004. Collaborative study of antibiotic medium 3 and flow cytometry for identification of amphotericin B-resistant Candida isolates. J. Clin. Microbiol. 42:2252–2254.

VOL. 24, 2011 29. Chaumentin, G., et al. 1996. Trichosporon inkin endocarditis: short-term evolution and clinical report. Clin. Infect. Dis. 23:396–397. 30. Chaves, G. M., M. A. Q. Cavalcanti, and A. L. F. Porto. 2003. Pathogenicity characteristics of stocked and fresh yeasts strains. Braz. J. Microbiol. 34: 197–202. 31. Chen, J., S. Shimura, K. Kirimura, and S. Usami. 1994. Purification of extracellular lipases from Trichosporon fermentans WU-C12. J. Ferment. Bioengin. 77:548–550. 32. Christakis, G., S. Perlorentzou, M. Aslanidou, A. Megalakaki, and A. Velegraki. 2005. Fatal Blastoschizomyces capitatus sepsis in a neutropenic patient with acute myeloid leukemia: first documented case from Greece. Mycoses 48:216–220. 33. Ciardo, D. E., G. Schar, E. C. Bottger, M. Altwegg, and P. P. Bosshard. 2006. Internal transcribed spacer sequencing versus biochemical profiling for identification of medically important yeasts. J. Clin. Microbiol. 44:77–84. 34. Colombo, A. L., et al. 1996. Gastrointestinal translocation as a possible source of candidemia in an AIDS patient. Rev. Inst. Med. Trop. Sao Paulo 38:197–200. 35. Cornely, O. A., K. Schmitz, and S. Aisenbrey. 2002. The first echinocandin: caspofungin. Mycoses 45:56–60. 36. Cuenca-Estrella, M., et al. 2003. Multicenter evaluation of the reproducibility of the proposed antifungal susceptibility testing method for fermentative yeasts of the Antifungal Susceptibility Testing Subcommittee of the European Committee on Antimicrobial Susceptibility Testing (AFST-EUCAST). Clin. Microbiol. Infect. 9:467–474. 37. Dag˘, A., and N. Cerikc¸iog˘lu. 2006. Investigation of some virulence factors of Trichosporon asahii strains isolated from the clinical samples of hospitalized patients. Mikrobiyol. Bul. 40:225–235. 38. De Hoog, G. S., M. T. Smith, and E. Gueho. 1986. A revision of the genus Geotrichum and its teleomorphs. Stud. Mycol. 29:1–131. 39. De Hoogs, G. S., J. Guarro, J. Gene, and M. J. Figueras. 2000. Atlas of clinical fungi, 2nd ed. Guanabara, Rio de Janeiro, Brazil. 40. De Pauw, B., et al. 2008. Revised definitions of invasive fungal disease from the European Organization for Research and Treatment of Cancer/Invasive Fungal Infections Cooperative Group and the National Institute of Allergy and Infectious Diseases Mycoses Study Group (EORTC/MSG) Consensus Group. Clin. Infect. Dis. 46:1813–1821. 41. Diaz, M. R., and J. W. Fell. 2004. High-throughput detection of pathogenic yeasts of the genus Trichosporon. J. Clin. Microbiol. 42:3696–3706. 42. Di Bonaventura, G., et al. 2006. Biofilm formation by the emerging fungal pathogen Trichosporon asahii: development, architecture, and antifungal resistance. Antimicrob. Agents Chemother. 50:3269–3276. 43. Dooley, D. P., M. L. Beckius, and B. S. Jeffrey. 1994. Misidentification of clinical yeast isolates by using the updated Vitek Yeast Biochemical Card. J. Clin. Microbiol. 32:2889–2892. 44. Ebright, J. R., M. R. Fairfax, and J. A. Vazquez. 2001. Trichosporon asahii, a non-Candida yeast that caused fatal septic shock in a patient without cancer or neutropenia. Clin. Infect. Dis. 33:E28–E30. 45. Ellner, K., M. E. McBride, T. Rosen, and D. Berman. 1991. Prevalence of Trichosporon beigelii. Colonization of normal perigenital skin. J. Med. Vet. Mycol. 29:99–103. 46. Erer, B., et al. 2000. Trichosporon beigelii: a life-threatening pathogen in immunocompromised hosts. Bone Marrow Transplant. 25:745–749. 47. Espinel-Ingroff, A. 1998. Comparison of in vitro activities of the new triazole SCH56592 and the echinocandins MK-0991 (L-743,872) and LY303366 against opportunistic filamentous and dimorphic fungi and yeasts. J. Clin. Microbiol. 36:2950–2956. 48. Espinel-Ingroff, A., et al. 1998. Comparison of RapID yeast plus system with API 20C system for identification of common, new, and emerging yeast pathogens. J. Clin. Microbiol. 36:883–886. 49. EUCAST. 2008. European Committee on Antimicrobial Susceptibility Testing. EUCAST Definitive Document EDef 7.1: method for the determination of broth dilution MICs of antifungal agents for fermentative yeasts. Clin. Microbiol. Infect. 14:398–405. 50. Febre, N., et al. 1999. Microbiological characteristics of yeasts isolated from urinary tracts of intensive care unit patients undergoing urinary catheterization. J. Clin. Microbiol. 37:1584–1586. 51. Fell, J. W., and G. Scorzetti. 2004. Reassignment of the basidiomycetous yeasts Trichosporon pullulans to Guehomyces pullulans gen. nov., comb. nov. and Hyalodendron lignicola to Trichosporon lignicola comb. nov. Int. J. Syst. Evol. Microbiol. 54:995–998. 52. Fenn, J., et al. 1994. Comparison of updated Vitek yeast biochemical card and API 20C yeast identification system. J. Clin. Microbiol. 32:1184–1187. 53. Fischman, O. 1973. Black piedra among Brazilian Indians. Rev. Inst. Med. Trop. Sao Paulo 15:103–106. 54. Fischman, O. 1965. Black piedra in Brazil. A contribution to its study in Manaus (state of Amazonas). Mycopathol. Mycol. Appl. 25:201–204. 55. Fleming, R. V., T. J. Walsh, and E. J. Anaissie. 2002. Emerging and less common fungal pathogens. Infect. Dis. Clin. North Am. 16:915–933. 56. Fonseca, F. L., et al. 2009. Structural and functional properties of the Trichosporon asahii glucuronoxylomannan. Fungal Genet. Biol. 46:496–505. 57. Fournier, S., et al. 2002. Use of voriconazole to successfully treat dissem-

TRICHOSPORON SPP. AND TRICHOSPORONOSIS

58.

59.

60. 61. 62.

63.

64.

65. 66. 67. 68.

69. 70.

71.

72.

73.

74. 75.

76.

77. 78.

79.

80.

81. 82.

83.

84.

85.

697

inated Trichosporon asahii infection in a patient with acute myeloid leukaemia. Eur. J. Clin. Microbiol. Infect. Dis. 21:892–896. Fox, A. 2006. Mass spectrometry for species or strain identification after culture or without culture: past, present, and future. J. Clin. Microbiol. 44:2677–2680. Fries, B. C., D. L. Goldman, R. Cherniak, R. Ju, and A. Casadevall. 1999. Phenotypic switching in Cryptococcus neoformans results in changes in cellular morphology and glucuronoxylomannan structure. Infect. Immun. 67: 6076–6083. Fuentefria, A. M., et al. 2008. Trichosporon insectorum sp. nov., a new anamorphic basidiomycetous killer yeast. Mycol. Res. 112:93–99. Ghannoum, M. A. 2000. Potential role of phospholipases in virulence and fungal pathogenesis. Clin. Microbiol. Rev. 13:122–143. Girmenia, C., et al. 2005. Invasive infections caused by Trichosporon species and Geotrichum capitatum in patients with hematological malignancies: a retrospective multicenter study from Italy and review of the literature. J. Clin. Microbiol. 43:1818–1828. Gold, I., B. Sommer, S. Urson, and M. Schewach-Millet. 1984. White piedra. A frequently misdiagnosed infection of hair. Int. J. Dermatol. 23: 621–623. Goodman, D., E. Pamer, A. Jakubowski, C. Morris, and K. Sepkowitz. 2002. Breakthrough trichosporonosis in a bone marrow transplant recipient receiving caspofungin acetate. Clin. Infect. Dis. 35:35–36. Groll, A. H., and T. J. Walsh. 2001. Uncommon opportunistic fungi: new nosocomial threats. Clin. Microbiol. Infect. 7(Suppl. 2):8–24. Gueho, E., G. S. de Hoog, and M. T. Smith. 1992. Neotypification of the genus Trichosporon. Antonie Van Leeuwenhoek 61:285–288. Gueho, E., L. Improvisi, G. S. de Hoog, and B. Dupont. 1994. Trichosporon on humans: a practical account. Mycoses 37:3–10. Gueho, E., M. T. Smith, and G. S. de Hoog. 1998. Trichosporon Behrend, p. 854–872. In C. P. Kurtzman and J. W. Fell (ed.), The yeasts, a taxonomic study, 4th ed. Elsevier, Amsterdam, The Netherlands. Gueho, E., et al. 1992. Contributions to a revision of the genus Trichosporon. Antonie Van Leeuwenhoek 61:289–316. Gujjari, P., S. O. Suh, K. Coumes, and J. J. Zhou. 10 May 2011, posting date. Characterization of oleaginous yeasts revealed two novel species: Trichosporon cacaoliposimilis sp. nov. and Trichosporon oleaginosus sp. nov. Mycologia doi:10.3852/10-403. Gujjari, P., S. O. Suh, C. F. Lee, and J. J. Zhou. 29 October 2010, posting date. Trichosporon xylopini sp. nov., a hemicellulose-degrading yeast isolated from wood-inhabiting beetle Xylopinus saperdioides. Int. J. Syst. Evol. Microbiol. doi:10.1099/ijs.0.028860-0. Gundes, S. G., S. Gulenc, and R. Bingol. 2001. Comparative performance of Fungichrom I, Candifast and API 20C Aux systems in the identification of clinically significant yeasts. J. Med. Microbiol. 50:1105–1110. Hajjeh, R. A., and H. M. Blumberg. 1995. Bloodstream infection due to Trichosporon beigelii in a burn patient: case report and review of therapy. Clin. Infect. Dis. 20:913–916. Hara, S., et al. 2007. Endophthalmitis due to Trichosporon beigelii in acute leukemia. Int. J. Hematol. 85:415–417. Hata, K., et al. 1996. In vitro and in vivo antifungal activities of ER-30346, a novel oral triazole with a broad antifungal spectrum. Antimicrob. Agents Chemother. 40:2237–2242. Haupt, H. M., W. G. Merz, W. E. Beschorner, W. P. Vaughan, and R. Saral. 1983. Colonization and infection with Trichosporon species in the immunosuppressed host. J. Infect. Dis. 147:199–203. Hogan, L. H., B. S. Klein, and S. M. Levitz. 1996. Virulence factors of medically important fungi. Clin. Microbiol. Rev. 9:469–488. Holland, S. M., Y. R. Shea, and J. Kwon-Chung. 2004. Regarding “Trichosporon pullulans infection in 2 patients with chronic granulomatous disease.” J. Allergy Clin. Immunol. 114:205–206. (Author reply, 114:206.) Hosoki, K., S. Iwamoto, T. Kumamoto, E. Azuma, and Y. Komada. 2008. Early detection of breakthrough trichosporonosis by serum PCR in a cord blood transplant recipient being prophylactically treated with voriconazole. J. Pediatr. Hematol. Oncol. 30:917–919. Hospenthal, D. R., C. K. Murray, and M. G. Rinaldi. 2004. The role of antifungal susceptibility testing in the therapy of candidiasis. Diagn. Microbiol. Infect. Dis. 48:153–160. Hoy, J., et al. 1986. Trichosporon beigelii infection: a review. Rev. Infect. Dis. 8:959–967. Hsiue, H. C., et al. 2010. Rapid identification of fungal pathogens in positive blood cultures using oligonucleotide array hybridization. Clin. Microbiol. Infect. 16:493–500. Huang, L. U., et al. 2001. A comparison of methods for yeast identification including CHROMagar Candida, Vitek system YBC and a traditional biochemical method. Zhonghua Yi Xue Za Zhi (Taipei) 64:568–574. Ichikawa, T., et al. 2004. Phenotypic switching and beta-N-acetylhexosaminidase activity of the pathogenic yeast Trichosporon asahii. Microbiol. Immunol. 48:237–242. Ikeda, R., M. Yokota, and T. Shinoda. 1996. Serological characterization of Trichosporon cutaneum and related species. Microbiol. Immunol. 40:813– 819.

698

COLOMBO ET AL.

86. Izumi, K., Y. Hisata, and S. Hazama. 2009. A rare case of infective endocarditis complicated by Trichosporon asahii fungemia treated by surgery. Ann. Thorac. Cardiovasc. Surg. 15:350–353. 87. Jain, N., F. Hasan, and B. C. Fries. 2008. Phenotypic switching in fungi. Curr. Fungal Infect. Rep. 2:180–188. 88. Jian, D. Y., W. C. Yang, T. W. Chen, and C. C. Lin. 2008. Trichosporon asahii following polymicrobial infection in peritoneal dialysis-associated peritonitis. Perit. Dial. Int. 28:100–101. 89. Kalkanci, A., et al. 2010. Molecular identification, genotyping, and drug susceptibility of the basidiomycetous yeast pathogen Trichosporon isolated from Turkish patients. Med. Mycol. 48:141–146. 90. Kaltreider, H. B. 1993. Hypersensitivity pneumonitis. West. J. Med. 159: 570–578. 91. Karashima, R., et al. 2002. Increased release of glucuronoxylomannan antigen and induced phenotypic changes in Trichosporon asahii by repeated passage in mice. J. Med. Microbiol. 51:423–432. 92. Kataoka-Nishimura, S., et al. 1998. Invasive infection due to Trichosporon cutaneum in patients with hematologic malignancies. Cancer 82:484–487. 93. Keay, S., D. W. Denning, and D. A. Stevens. 1991. Endocarditis due to Trichosporon beigelii: in vitro susceptibility of isolates and review. Rev. Infect. Dis. 13:383–386. 94. Kemker, B. J., P. F. Lehmann, J. W. Lee, and T. J. Walsh. 1991. Distinction of deep versus superficial clinical and nonclinical isolates of Trichosporon beigelii by isoenzymes and restriction fragment length polymorphisms of rDNA generated by polymerase chain reaction. J. Clin. Microbiol. 29:1677–1683. 95. Khanna, R., D. G. Oreopoulos, S. Vas, D. McNeely, and W. McCready. 1980. Treating fungal infections. Br. Med. J. 280:1147–1148. 96. Kiken, D. A., et al. 2006. White piedra in children. J. Am. Acad. Dermatol. 55:956–961. 97. Kitch, T. T., M. R. Jacobs, M. R. McGinnis, and P. C. Appelbaum. 1996. Ability of RapID Yeast Plus system to identify 304 clinically significant yeasts within 5 hours. J. Clin. Microbiol. 34:1069–1071. 98. Kontoyiannis, D. P., et al. 2004. Trichosporonosis in a tertiary care cancer center: risk factors, changing spectrum and determinants of outcome. Scand. J. Infect. Dis. 36:564–569. 99. Krcmery, V., Jr., et al. 1999. Hematogenous trichosporonosis in cancer patients: report of 12 cases including 5 during prophylaxis with itraconazol. Support Care Cancer 7:39–43. 100. Kumar, S. S., L. Kumar, V. Sahai, and R. Gupta. 2009. A thiol-activated lipase from Trichosporon asahii MSR 54: detergent compatibility and presoak formulation for oil removal from soiled cloth at ambient temperature. J. Ind. Microbiol. Biotechnol. 36:427–432. 101. Kwon-Chung, K. J., and J. E. Bennett. 1992. Medical mycology. Lea & Febiger, Philadelphia, PA. 102. Land, G. A., I. F. Salkin, M. el-Zaatari, M. R. McGinnis, and G. Hashem. 1991. Evaluation of the Baxter-MicroScan 4-hour enzyme-based yeast identification system. J. Clin. Microbiol. 29:718–722. 103. Landlinger, C., et al. 2009. Species-specific identification of a wide range of clinically relevant fungal pathogens by use of Luminex xMAP technology. J. Clin. Microbiol. 47:1063–1073. 104. Leaw, S. N., H. C. Chang, R. Barton, J. P. Bouchara, and T. C. Chang. 2007. Identification of medically important Candida and non-Candida yeast species by an oligonucleotide array. J. Clin. Microbiol. 45:2220–2229. 105. Leaw, S. N., et al. 2006. Identification of medically important yeast species by sequence analysis of the internal transcribed spacer regions. J. Clin. Microbiol. 44:693–699. 106. Li, H., Q. Lu, Z. Wan, and J. Zhang. 2010. In vitro combined activity of amphotericin B, caspofungin and voriconazole against clinical isolates of Trichosporon asahii. Int. J. Antimicrob. Agents 35:550–552. 107. Lopes, J. O., S. H. Alves, C. Klock, L. T. Oliveira, and N. R. Dal Forno. 1997. Trichosporon inkin peritonitis during continuous ambulatory peritoneal dialysis with bibliography review. Mycopathologia 139:15–18. 108. Lussier, N., M. Laverdiere, J. Delorme, K. Weiss, and R. Dandavino. 2000. Trichosporon beigelii funguria in renal transplant recipients. Clin. Infect. Dis. 31:1299–1301. 109. Lyman, C. A., et al. 1995. Detection and quantitation of the glucuronoxylomannan-like polysaccharide antigen from clinical and nonclinical isolates of Trichosporon beigelii and implications for pathogenicity. J. Clin. Microbiol. 33:126–130. 110. Macedo, D. P., et al. 2011. Trichosporon inkin esophagitis: an uncommon disease in a patient with pulmonary cancer. Mycopathologia 171:279–283. 111. Madariaga, M. G., A. Tenorio, and L. Proia. 2003. Trichosporon inkin peritonitis treated with caspofungin. J. Clin. Microbiol. 41:5827–5829. 112. Makino, H., J. Fujimoto, and K. Watanabe. 2010. Development and evaluation of a real-time quantitative PCR assay for detection and enumeration of yeasts of public health interest in dairy products. Int. J. Food Microbiol. 140:76–83. 113. Manzella, J. P., I. J. Berman, and M. D. Kukrika. 1982. Trichosporon beigelii fungemia and cutaneous dissemination. Arch. Dermatol. 118:343– 345. 114. Marier, R., et al. 1978. Trichosporon cutaneum endocarditis. Scand. J. Infect. Dis. 10:225–226.

CLIN. MICROBIOL. REV. 115. Marklein, G., et al. 2009. Matrix-assisted laser desorption ionization–time of flight mass spectrometry for fast and reliable identification of clinical yeast isolates. J. Clin. Microbiol. 47:2912–2917. 116. Martinez-Lacasa, J., et al. 1991. Long-term survival of a patient with prosthetic valve endocarditis due to Trichosporon beigelii. Eur. J. Clin. Microbiol. Infect. Dis. 10:756–758. 117. Marty, F. M., D. H. Barouch, E. P. Coakley, and L. R. Baden. 2003. Disseminated trichosporonosis caused by Trichosporon loubieri. J. Clin. Microbiol. 41:5317–5320. 118. Matsue, K., H. Uryu, M. Koseki, N. Asada, and M. Takeuchi. 2006. Breakthrough trichosporonosis in patients with hematologic malignancies receiving micafungin. Clin. Infect. Dis. 42:753–757. 119. Mekha, N., et al. 2010. Genotyping and antifungal drug susceptibility of the pathogenic yeast Trichosporon asahii isolated from Thai patients. Mycopathologia 169:67–70. 120. Melcher, G. P., et al. 1991. Demonstration of a cell wall antigen crossreacting with cryptococcal polysaccharide in experimental disseminated trichosporonosis. J. Clin. Microbiol. 29:192–196. 121. Mendez-Tovar, L. J., et al. 2006. Micosis among five highly underprivileged Mexican communities. Gaceta Med. Mexico 142:381–386. 122. Meyer, M. H., et al. 2002. Chronic disseminated Trichosporon asahii infection in a leukemic child. Clin. Infect. Dis. 35:22–25. 123. Middelhoven, W. J., G. Scorzetti, and J. W. Fell. 2004. Systematics of the anamorphic basidiomycetous yeast genus Trichosporon Behrend with the description of five novel species: Trichosporon vadense, T. smithiae, T. dehoogii, T. scarabaeorum and T. gamsii. Int. J. Syst. Evol. Microbiol. 54:975–986. 124. Mizobe, T., M. Ando, H. Yamasaki, K. Onoue, and A. Misaki. 1995. Purification and characterization of the serotype-specific polysaccharide antigen of Trichosporon cutaneum serotype II: a disease-related antigen of Japanese summer-type hypersensitivity pneumonitis. Clin. Exp. Allergy 25:265–272. 125. Molnar, O., G. Schatzmayr, E. Fuchs, and H. Prillinger. 2004. Trichosporon mycotoxinivorans sp. nov., a new yeast species useful in biological detoxification of various mycotoxins. Syst. Appl. Microbiol. 27:661–671. 126. Moretti-Branchini, M. L., et al. 2001. Trichosporon species infection in bone marrow transplanted patients. Diagn. Microbiol. Infect. Dis. 39:161–164. 127. Mori, Y., et al. 2005. Marked renal damage in a child with hydronephrosis infected by Trichosporon asahii. Pediatr. Nephrol. 20:234–236. 128. Moylett, E. H., J. Chinen, and W. T. Shearer. 2003. Trichosporon pullulans infection in 2 patients with chronic granulomatous disease: an emerging pathogen and review of the literature. J. Allergy Clin. Immunol. 111:1370–1374. 129. Nagai, H., Y. Yamakami, A. Hashimoto, I. Tokimatsu, and M. Nasu. 1999. PCR detection of DNA specific for Trichosporon species in serum of patients with disseminated trichosporonosis. J. Clin. Microbiol. 37:694–699. 130. Nagano, Y., et al. 2010. Comparison of techniques to examine the diversity of fungi in adult patients with cystic fibrosis. Med. Mycol. 48:166–176. 131. Nakajima, M., T. Sugita, and Y. Mikami. 2007. Granuloma associated with Trichosporon asahii infection in the lung: unusual pathological findings and PCR detection of Trichosporon DNA. Med. Mycol. 45:641–644. 132. NCCLS/CLSI. 2008. Reference method for broth dilution antifungal susceptibility testing of yeasts. M27-A3, vol. 28. NCCLS/CLSI, Wayne, PA. 133. Nishiura, Y., et al. 1997. Assignment and serotyping of Trichosporon species: the causative agents of summer-type hypersensitivity pneumonitis. J. Med. Vet. Mycol. 35:45–52. 134. Nucci, M., and E. Anaissie. 2001. Revisiting the source of candidemia: skin or gut? Clin. Infect. Dis. 33:1959–1967. 135. Obana, Y., M. Sano, T. Jike, T. Homma, and N. Nemoto. 2010. Differential diagnosis of trichosporonosis using conventional histopathological stains and electron microscopy. Histopathology 56:372–383. 136. Odabasi, Z., et al. 2004. Beta-D-glucan as a diagnostic adjunct for invasive fungal infections: validation, cutoff development, and performance in patients with acute myelogenous leukemia and myelodysplastic syndrome. Clin. Infect. Dis. 39:199–205. 137. Padhye, A. A., et al. 2003. Trichosporon loubieri infection in a patient with adult polycystic kidney disease. J. Clin. Microbiol. 41:479–482. 138. Pagano, L., et al. 2006. The epidemiology of fungal infections in patients with hematologic malignancies: the SEIFEM-2004 study. Haematologicaae 91:1068–1075. 139. Pagnocca, F. C., et al. 2010. Yeasts isolated from a fungus-growing ant nest, including the description of Trichosporon chiarellii sp. nov., an anamorphic basidiomycetous yeast. Int. J. Syst. Evol. Microbiol. 60:1454–1459. 140. Paphitou, N. I., et al. 2002. In vitro antifungal susceptibilities of Trichosporon species. Antimicrob. Agents Chemother. 46:1144–1146. 141. Perparim, K., et al. 1996. In vitro susceptibility of Trichosporon beigelii to antifungal agents. J. Chemother. 8:445–448. 142. Pfaller, M. A. 1995. Epidemiology of fungal infections: the promise of molecular typing. Clin. Infect. Dis. 20:1535–1539. 143. Pfaller, M. A., and D. J. Diekema. 2004. Rare and emerging opportunistic fungal pathogens: concern for resistance beyond Candida albicans and Aspergillus fumigatus. J. Clin. Microbiol. 42:4419–4431. 144. Pincus, D. H., S. Orenga, and S. Chatellier. 2007. Yeast identification— past, present, and future methods. Med. Mycol. 45:97–121. 145. Ramage, G., E. Mowat, B. Jones, C. Williams, and J. Lopez-Ribot. 2009.

VOL. 24, 2011

146.

147.

148.

149.

150.

151.

152.

153. 154.

155.

156. 157. 158.

159. 160.

161.

162.

163.

164.

165.

166.

167.

168. 169.

170.

171.

Our current understanding of fungal biofilms. Crit. Rev. Microbiol. 35:340– 345. Ramani, R., S. Gromadzki, D. H. Pincus, I. F. Salkin, and V. Chaturvedi. 1998. Efficacy of API 20C and ID 32C systems for identification of common and rare clinical yeast isolates. J. Clin. Microbiol. 36:3396–3398. Ramos, J. M., M. Cuenca-Estrella, F. Gutierrez, M. Elia, and J. L. Rodriguez-Tudela. 2004. Clinical case of endocarditis due to Trichosporon inkin and antifungal susceptibility profile of the organism. J. Clin. Microbiol. 42:2341–2344. Reinhart, H. H., D. M. Urbanski, S. D. Harrington, and J. D. Sobel. 1988. Prosthetic valve endocarditis caused by Trichosporon beigelii. Am. J. Med. 84:355–358. Reyes, C. V., M. M. Stanley, and J. W. Rippon. 1985. Trichosporon beigelii endocarditis as a complication of peritoneovenous shunt. Hum. Pathol. 16:857–859. Rodrigues, M. L., F. L. Fonseca, S. Frases, A. Casadevall, and L. Nimrichter. 2009. The still obscure attributes of cryptococcal glucuronoxylomannan. Med. Mycol. 47:783–788. Rodriguez-Tudela, J. L., et al. 2005. Susceptibility patterns and molecular identification of Trichosporon species. Antimicrob. Agents Chemother. 49: 4026–4034. Rodriguez-Tudela, J. L., et al. 2007. Genotype distribution of clinical isolates of Trichosporon asahii based on sequencing of intergenic spacer 1. Diagn. Microbiol. Infect. Dis. 58:435–440. Roshan, A. S., C. Janaki, and B. Parveen. 2009. White piedra in a mother and daughter. Int. J. Trichol. 1:140–141. Ruan, S. Y., J. Y. Chien, and P. R. Hsueh. 2009. Invasive trichosporonosis caused by Trichosporon asahii and other unusual Trichosporon species at a medical center in Taiwan. Clin. Infect. Dis. 49:11–17. Ruiz-Esmenjaud, J., R. Arenas, M. Rodriguez-Alvarez, E. Monroy, and R. Felipe Fernandez. 2003. Tinea pedis and onychomycosis in children of the Mazahua Indian community in Mexico. Gaceta Med. Mexico 139:215–220. Salazar, G. E., and J. R. Campbell. 2002. Trichosporonosis, an unusual fungal infection in neonates. Pediatr. Infect. Dis. J. 21:161–165. Salkin, I. F., M. A. Gordon, W. A. Samsonoff, and C. L. Rieder. 1985. Blastoschizomyces capitatus, a new combination. Mycotaxon 22:375–380. Sano, M., et al. 2007. Supplemental utility of nested PCR for the pathological diagnosis of disseminated trichosporonosis. Virchows Arch. 451: 929–935. Schwartz, R. A. 2004. Superficial fungal infections. Lancet 364:1173–1182. Serena, C., F. Gilgado, M. Marine, F. J. Pastor, and J. Guarro. 2006. Efficacy of voriconazole in a guinea pig model of invasive trichosporonosis. Antimicrob. Agents Chemother. 50:2240–2243. Serena, C., F. J. Pastor, F. Gilgado, E. Mayayo, and J. Guarro. 2005. Efficacy of micafungin in combination with other drugs in a murine model of disseminated trichosporonosis. Antimicrob. Agents Chemother. 49:497– 502. Shang, S. T., Y. S. Yang, and M. Y. Peng. 2010. Nosocomial Trichosporon asahii fungemia in a patient with secondary hemochromatosis: a rare case report. J. Microbiol. Immunol. Infect. 43:77–80. Shimazu, K., M. Ando, T. Sakata, K. Yoshida, and S. Araki. 1984. Hypersensitivity pneumonitis induced by Trichosporon cutaneum. Am. Rev. Respir. Dis. 130:407–411. Sidarous, M. G., M. V. O’Reilly, and C. E. Cherubin. 1994. A case of Trichosporon beigelii endocarditis 8 years after aortic valve replacement. Clin. Cardiol. 17:215–219. Silvestre, A. M., Jr., M. A. R. Miranda, and Z. P. Camargo. 2010. Trichosporon species isolated from the perigenital region, urine and catheters of a Brazilian population. Braz. J. Microbiol. 41:628–634. Spiess, B., et al. 2007. DNA microarray-based detection and identification of fungal pathogens in clinical samples from neutropenic patients. J. Clin. Microbiol. 45:3743–3753. Stevenson, L. G., S. K. Drake, Y. R. Shea, A. M. Zelazny, and P. R. Murray. 2010. Evaluation of matrix-assisted laser desorption ionization–time of flight mass spectrometry for identification of clinically important yeast species. J. Clin. Microbiol. 48:3482–3486. St. Germain, G., and D. Beauchesne. 1991. Evaluation of the MicroScan Rapid Yeast Identification panel. J. Clin. Microbiol. 29:2296–2299. Sugita, T., R. Ikeda, and A. Nishikawa. 2004. Analysis of Trichosporon isolates obtained from the houses of patients with summer-type hypersensitivity pneumonitis. J. Clin. Microbiol. 42:5467–5471. Sugita, T., et al. 2005. Trichosporon species isolated from guano samples obtained from bat-inhabited caves in Japan. Appl. Environ. Microbiol. 71:7626–7629. Sugita, T., et al. 1997. Partial sequences of large subunit ribosomal DNA of a new yeast species, Trichosporon domesticum and related species. Microbiol. Immunol. 41:571–573.

TRICHOSPORON SPP. AND TRICHOSPORONOSIS

699

172. Sugita, T., M. Nakajima, R. Ikeda, T. Matsushima, and T. Shinoda. 2002. Sequence analysis of the ribosomal DNA intergenic spacer 1 regions of Trichosporon species. J. Clin. Microbiol. 40:1826–1830. 173. Sugita, T., et al. 2001. A nested PCR assay to detect DNA in sera for the diagnosis of deep-seated trichosporonosis. Microbiol. Immunol. 45:143– 148. 174. Sugita, T., A. Nishikawa, R. Ikeda, and T. Shinoda. 1999. Identification of medically relevant Trichosporon species based on sequences of internal transcribed spacer regions and construction of a database for Trichosporon identification. J. Clin. Microbiol. 37:1985–1993. 175. Sugita, T., A. Nishikawa, and T. Shinoda. 1998. Rapid detection of species of the opportunistic yeast Trichosporon by PCR. J. Clin. Microbiol. 36: 1458–1460. 176. Sugita, T., A. Nishikawa, and T. Shinoda. 1994. Reclassification of Trichosporon cutaneum by DNA relatedness by using the spectrophotometric method and chemiluminometric method. J. Gen. Appl. Microbiol. 40:397– 408. 177. Sugita, T., A. Nishikawa, T. Shinoda, and H. Kume. 1995. Taxonomic position of deep-seated, mucosa-associated, and superficial isolates of Trichosporon cutaneum from trichosporonosis patients. J. Clin. Microbiol. 33:1368–1370. 178. Surmont, I., et al. 1990. First report of chronic meningitis caused by Trichosporon beigelii. Eur. J. Clin. Microbiol. Infect. Dis. 9:226–229. 179. Suzuki, K., et al. 2010. Fatal Trichosporon fungemia in patients with hematologic malignancies. Eur. J. Haematol. 84:441–447. 180. Taj-Aldeen, S. J., et al. 2009. Molecular identification and susceptibility of Trichosporon species isolated from clinical specimens in Qatar: isolation of Trichosporon dohaense Taj-Aldeen, Meis & Boekhout sp. nov. J. Clin. Microbiol. 47:1791–1799. 181. Therizol-Ferly, M., et al. 1994. White piedra and Trichosporon species in equatorial Africa. II. Clinical and mycological associations: an analysis of 449 superficial inguinal specimens. Mycoses 37:255–260. 182. Thomas, D., A. Mogahed, J. P. Leclerc, and Y. Grosgogeat. 1984. Prosthetic valve endocarditis caused by Trichosporon cutaneum. Int. J. Cardiol. 5:83–87. 183. Tokimatsu, I., et al. 2007. The prophylactic effectiveness of various antifungal agents against the progression of trichosporonosis fungemia to disseminated disease in a neutropenic mouse model. Int. J. Antimicrob. Agents. 29:84–88. 184. Trentin, L., et al. 1990. Mechanisms accounting for lymphocytic alveolitis in hypersensitivity pneumonitis. J. Immunol. 145:2147–2154. 185. Uzun, O., S. Arikan, S. Kocagoz, B. Sancak, and S. Unal. 2000. Susceptibility testing of voriconazole, fluconazole, itraconazole and amphotericin B against yeast isolates in a Turkish University Hospital and effect of time of reading. Diagn. Microbiol. Infect. Dis. 38:101–107. 186. Walsh, T. J. 1990. Role of surveillance cultures in prevention and treatment of fungal infections. NCI Monogr. 9:43–45. 187. Walsh, T. J. 1989. Trichosporonosis. Infect. Dis. Clin. North Am. 3:43–52. 188. Walsh, T. J., et al. 2004. Infections due to emerging and uncommon medically important fungal pathogens. Clin. Microbiol. Infect. 10:44–66. 189. Walsh, T. J., et al. 1992. Experimental Trichosporon infection in persistently granulocytopenic rabbits: implications for pathogenesis, diagnosis, and treatment of an emerging opportunistic mycosis. J. Infect. Dis. 166:121–133. 190. Walsh, T. J., et al. 1990. Trichosporon beigelii, an emerging pathogen resistant to amphotericin B. J. Clin. Microbiol. 28:1616–1622. 190a.Watson, K. C., and S. Kallichurum. 1970. Brain abscess due to Trichosporon cutaneum. J. Med. Microbiol. 3:191–193. 191. Wolf, D. G., et al. 2001. Multidrug-resistant Trichosporon asahii infection of nongranulocytopenic patients in three intensive care units. J. Clin. Microbiol. 39:4420–4425. 192. Yamada, Y., E. Nakazawa, and K. Kondo. 1982. The coenzyme Q system in strains of Trichosporon species and related organisms. J. Gen. Appl. Microbiol. 28:355–358. 193. Yamamoto, K., et al. 1997. Experimental disseminated trichosporonosis in mice: tissue distribution and therapy with antifungal agents. J. Med. Vet. Mycol. 35:411–418. 194. Yoo, C. G., et al. 1997. Summer-type hypersensitivity pneumonitis outside Japan: a case report and the state of the art. Respirology 2:75–77. 195. Yoshida, K., M. Ando, T. Sakata, and S. Araki. 1988. Environmental mycological studies on the causative agent of summer-type hypersensitivity pneumonitis. J. Allergy Clin. Immunol. 81:475–483. 196. Yoshida, K., M. Ando, T. Sakata, and S. Araki. 1989. Prevention of summer-type hypersensitivity pneumonitis: effect of elimination of Trichosporon cutaneum from the patients’ homes. Arch. Environ. Health 44:317–322. 197. Youker, S. R., and R. J. Andreozzi. 2003. White piedra: further evidence of a synergistic infection. J. Am. Acad. Dermatol. 49:746–749.

Continued next page

700

COLOMBO ET AL.

Arnaldo Lopes Colombo is a Professor of Medicine at the Division of Infectious Diseases of the Federal University of Sa˜o Paulo-UNIFESP, Brazil. He is currently the Dean of Research of the Federal University of Sa˜o Paulo and Head of the Special Mycology Laboratory at the same university, a reference laboratory in Brazil and Latin America for yeast identification, typing, and antifungal susceptibility testing. Dr. Colombo obtained his M.D. from UNIFESP in 1983 and continued his residency training in internal medicine and infectious diseases there. He completed his fellowship training in medical mycology at the University of Texas Health Science Center at San Antonio, TX. He received his Ph.D. degree from UNIFESP in 1994. He has organized several multicenter surveillance studies to characterize the epidemiology of Candida and has actively participated in several surveillance programs to evaluate emergent fungal pathogens and antifungal resistance. Ana Carolina Barbosa Padovan is a biologist and received her Ph.D. from the Department of Microbiology, Immunology and Parasitology of the Federal University of Sa˜o Paulo-UNIFESP, Brazil, on the evolution of the Ascomycota fungi and the study of Candida albicans adherence genes and biofilm formation. She received postdoctoral training at Professor Roberto Kolter’s lab in the Department of Microbiology and Molecular Genetics at Harvard Medical School, Boston, MA, on the study of microbial interactions that inhibit the expression of Candida albicans virulence factors. Currently, she is a postdoctoral fellow at the Special Mycology Laboratory-UNIFESP, and her interests involve the development and application of molecular diagnostics of emergent fungal pathogens and the interactions between the normal human microbiota and fungi in order to prevent superficial and invasive fungal infections.

CLIN. MICROBIOL. REV. Guilherme Maranha ˜o Chaves is currently a Lecturer on Basic and Clinical Microbiology at the Department of Clinical and Toxicological Analysis at the Federal University of Rio Grande do Norte, UFRN, Brazil. He obtained his Ph.D. in molecular microbiology from University of Aberdeen, under the supervision of Professor Frank C. Odds, on the subject of oxidative stress and virulence in Candida albicans. He next received postdoctoral training at the Special Mycology Laboratory-UNIFESP on the epidemiological and genetic characterization of Trichosporon and Candida clinical isolates. His research interests are in molecular identification and genotyping and the characterization of virulence factors of Candida and emergent yeast pathogens.

CLINICAL MICROBIOLOGY REVIEWS, Oct. 2011, p. 701–717 0893-8512/11/$12.00 doi:10.1128/CMR.00020-11 Copyright © 2011, American Society for Microbiology. All Rights Reserved.

Vol. 24, No. 4

Clinical Manifestations, Diagnosis, and Treatment of Mycobacterium haemophilum Infections Jerome A. Lindeboom,1 Lesla E. S. Bruijnesteijn van Coppenraet,2 Dick van Soolingen,3,4 Jan M. Prins,5 and Eduard J. Kuijper6* Department of Oral and Maxillofacial Surgery, Academic Medical Center, Amsterdam, The Netherlands1; Department of Medical Microbiology and Infectious Diseases, Isala Clinics, Zwolle, The Netherlands2; Mycobacterial Reference Laboratory, National Institute for Public Health and the Environment, Bilthoven, The Netherlands3; Department of Pulmonary Diseases and Department of Microbiology, Radboud University Nijmegen Medical Centre, P.O. Box 9101, 6500HB Nijmegen, The Netherlands4; Department of Internal Medicine, Division of Infectious Diseases, Tropical Medicine and AIDS, Academic Medical Center, Amsterdam, The Netherlands5; and Department of Medical Microbiology, Center of Infectious Diseases, Leiden University Medical Center, Leiden, The Netherlands6 INTRODUCTION .......................................................................................................................................................702 GENERAL DESCRIPTION AND TAXONOMY ....................................................................................................702 CLINICAL PRESENTATION ...................................................................................................................................702 IMMUNOCOMPROMISED PATIENTS ................................................................................................................703 Cutaneous Manifestations .....................................................................................................................................703 Pyomyositis ..............................................................................................................................................................703 Disseminated and Pulmonary Infections.............................................................................................................703 Ophthalmologic Manifestations............................................................................................................................705 Osteomyelitis ...........................................................................................................................................................705 Uncommon Clinical Presentations .......................................................................................................................705 Central catheter infections ................................................................................................................................705 Epididymal abscess.............................................................................................................................................706 Mixed infections..................................................................................................................................................706 IMMUNOCOMPETENT PATIENTS ......................................................................................................................706 Adult Infections.......................................................................................................................................................706 Cervicofacial infections ......................................................................................................................................706 “Other” skin lesions...........................................................................................................................................706 Pediatric M. haemophilum Infections ...................................................................................................................708 Cervicofacial infections ......................................................................................................................................708 Inguinal lymphadenitis ......................................................................................................................................708 Pulmonary involvement......................................................................................................................................708 ANIMAL INFECTIONS.............................................................................................................................................708 PATHOGENESIS........................................................................................................................................................709 EPIDEMIOLOGY .......................................................................................................................................................709 Typing of M. haemophilum.....................................................................................................................................709 Environmental Findings.........................................................................................................................................709 DIAGNOSTICS ...........................................................................................................................................................710 Skin Testing.............................................................................................................................................................710 Histopathology.........................................................................................................................................................710 Microscopy ...............................................................................................................................................................710 Culture......................................................................................................................................................................710 Molecular Identification Methods ........................................................................................................................711 Direct Detection Methods......................................................................................................................................711 Diagnostic Approach ..............................................................................................................................................711 ANTIMICROBIAL SUSCEPTIBILITY....................................................................................................................712 TREATMENT..............................................................................................................................................................712 Immunocompromised Patients .............................................................................................................................713 Skin lesions..........................................................................................................................................................713 Disseminated infection/pulmonary infection...................................................................................................713 Pyomyositis ..........................................................................................................................................................713

* Corresponding author. Mailing address: Department of Medical Microbiology, Center of Infectious Diseases, Leiden University Medical Center, P.O. Box 9600, Leiden 2300 RC, The Netherlands. Phone: 071 5269111. Fax: 071 5248148. E-mail: [email protected]. 701

702

LINDEBOOM ET AL.

CLIN. MICROBIOL. REV.

Skeletal infections/osteomyelitis .......................................................................................................................713 Immunocompetent Patients...................................................................................................................................713 Immunocompetent adults ..................................................................................................................................713 Immunocompetent children...............................................................................................................................713 Treatment Outcome................................................................................................................................................713 RECOMMENDATIONS AND CONCLUSION ......................................................................................................714 REFERENCES ............................................................................................................................................................714 INTRODUCTION Mycobacterium haemophilum is an acid-fast bacillus (AFB) belonging to the group of nontuberculous mycobacteria (NTM) frequently found in environmental habitats, which can colonize and occasionally infect humans and animals (98). M. haemophilum can cause localized or disseminated disease in immunocompromised hosts and is a rare cause of disease in immunologically competent individuals. In 1996, Saubolle et al. (123) presented an overview of 64 cases reported in the literature. Since that time, another 154 cases have been reported. The purpose of this review is to present an update of the clinical picture, diagnostic approach, and therapeutic options for M. haemophilum infections. GENERAL DESCRIPTION AND TAXONOMY M. haemophilum, or the “blood-loving” mycobacterium, is a slowly growing AFB that differs from all other identified Mycobacterium species in preferring a lower growth temperature and having a unique culture requirement for iron supplementation. Thus, the classification of mycobacteria into several Runyon groups based on growth characteristics and pigment production may not be applicable to M. haemophilum. Many infections with M. haemophilum likely remain unrecognized, although suspicion should arise when AFB are visualized in smears and when cultures fail to yield an etiologic agent. M. haemophilum was first described in 1978 as a pathogen causing skin infections most frequently in immunocompromised patients, which may explain its preferred growth temperature of 30°C (130). In 1981, Dawson and colleagues described a case of submandibular lymphadenitis due to M. haemophilum in an otherwise healthy child (31), and M. haemophilum has since been recognized as an emerging pathogen in a variety of syndromes. The microorganism is now also known to cause cutaneous and subcutaneous infections, septic arthritis, osteomyelitis, and pneumonitis in immunocompromised patients. Cervicofacial lymphadenitis is the most common manifestation in immunocompetent children. Reports of such cases originate from all continents. However, although our understanding of M. haemophilum infections in humans has increased considerably in recent years, the natural habitat and how an infection is acquired remain unknown. M. haemophilum most resembles Mycobacterium marinum and M. ulcerans in regard to its role in skin infections. The relatedness can also be observed for genomic traits, as all three species have a relatively low GC content compared to those of most other Mycobacterium species. Some interesting similarities also exist between M. haemophilum and Mycobacterium leprae. First, the fatty acid docosanoic acid is found in abundant quantities in both species. Second, M. haemophilum has also been shown to possess a

specific phenolic glycolipid antigen that closely resembles the corresponding lipid in M. leprae (10). Third, M. leprae has major membrane protein I (35 kDa), which is absent in members of the M. tuberculosis complex, but homologous sequences have been detected in M. avium, M. haemophilum, and M. smegmatis (159). Taxonomic relationships between mycobacteria can be investigated by comparing the sequences of gene targets used to differentiate species, such as ribosomal gene fragments (i.e., the 16S rRNA gene and internal transcribed spacer [ITS]) and housekeeping genes (i.e., hsp65 and rpoB). The taxonomic relationship between M. haemophilum and other Mycobacterium species is not completely clear because different panels of mycobacterial species were included in previous studies, and different gene fragments were used in alignments: the 16S rRNA, rpoB, hsp65, and sod genes. A phylogenetic analysis of 500-bp 5⬘ 16S rRNA gene sequences in the RIDOM database indicated that M. leprae, M. malmoense, and M. bohemicum are the species genetically most closely related to M. haemophilum (62). Another tree constructed from the 16S rRNA gene sequences from 80 species indicated that M. leprae and the M. avium complex are closely related (50). A study using an unrooted phylogenetic analysis of 16S rRNA gene sequences (1,325 bp) from 18 species showed that M. bohemicum and M. szulgai are the most closely related species (58), and M. leprae has a relatively large genetic distance from M. haemophilum. Last, using a multigene approach, including the sod, 16S rRNA, hsp65, and rpoB genes, Devulder and colleagues showed that M. haemophilum has no immediate neighboring species, although M. leprae was not included in this analysis (34). At the National Institute for Public Health and the Environment (RIVM), about 700 nucleotides of the 3⬘ end of the rpoB gene were sequenced. With the sequence data obtained, a dendrogram (Fig. 1) was created by using BioNumerics software (Applied Maths, Kortrijk, Belgium). Based on rpoB similarities, the closest relationship was observed for M. leprae (93.5%). Other mycobacterial species were at larger genetic distances, with M. gordonae (92%), M. malmoense (91.7%), M. avium (91%), and M. szulgai being most closely related (R. de Zwaan, RIVM, unpublished data). CLINICAL PRESENTATION In contrast to infections caused by M. tuberculosis, M. haemophilum is not a reportable infection, and the number of cases may be higher than what is represented by published case reports. A second reason for the underestimation of the actual number of M. haemophilum infections is the difficulties in diagnosing the disease. Based on the available literature, two groups appear to be at risk for M. haemophilum infection (123). The main group consists of severely immunocompromised patients, in whom M. haemophilum occurs as an oppor-

VOL. 24, 2011

MYCOBACTERIUM HAEMOPHILUM INFECTIONS

703

(47). Skin lesions typically evolve from papules to asymptomatic pustules and eventually to very painful deep-seated ulcers. The erythematous or violaceous papules and/or nodules are usually painless at first, but they can develop into potentially very painful abscesses or ulcers. Patients with cutaneous and articular manifestations have a more favorable prognosis than those with pulmonary involvement (126). An overview of the skin infections reported since the review by Saubolle et al. (123) is presented in Table 1. Thirty-three new cases have been reported, with a median age of the patients of 48 years (range, 14 months to 67 years). The sex distribution was equal. Most of the reports were from the United States (10 cases), followed by Germany (4 cases), Australia (4 cases), and Singapore (4 cases). The majority of the patients had a history of solid-organ transplant or AIDS. Cutaneous lesions have been rarely reported for children (21, 28), but the manifestations of the skin lesions are similar to those of immunosuppressed adults. Pyomyositis

FIG. 1. Dendrogram made by using the rpoB gene sequences of 29 mycobacterial species. M. leprae was most closely associated (93.5%).

tunistic infection (1, 2). M. haemophilum is being increasingly recognized in persons who are severely immunocompromised by HIV infection; after renal, bone marrow, or cardiac transplantation; or after treatment for lymphoma or rheumatoid arthritis. The second at-risk group is otherwise healthy children, who typically develop cervical and perihilar lymphadenitis similar to that caused by infection with the Mycobacterium avium complex (3, 90, 164).

Mycobacterial infection of the skeletal muscle is very rare; in particular, large muscles are involved, and the condition usually presents as localized muscle involvement through direct extension from a proximal focus of infection. Only four cases of pyomyositis caused by M. haemophilum have been reported (70, 82, 124, 127). In a recent report by Lee et al. (82), a 23-year-old immunosuppressed female patient with multiple, tender, erythematous, and palpable fluctuant abscesses on the left leg due to an M. haemophilum infection was described. In another case, the patient had been on long-term steroid treatment for polymyositis and presented with ulcerations over both thighs and the left arm after a year of steroid therapy (127). A 24-year-old female renal transplant recipient was described as having tender, erythematous, and palpable fluctuant swelling on the left calf (70). The patient had undergone kidney transplantation 8 years earlier, after which she had been on immunosuppressive treatment with cyclosporine and mycophenolate mofetil. Disseminated and Pulmonary Infections

IMMUNOCOMPROMISED PATIENTS Cutaneous Manifestations M. haemophilum causes mainly skin lesions in immunocompromised patients (42, 150). Cutaneous infections with potentially pathogenic mycobacterial species are important for the differential diagnosis of skin lesions in these patients (36, 61, 93, 106). M. haemophilum infections have been reported, especially in patients with lymphoma or HIV and in organ transplant recipients (19, 66, 81, 84, 112, 153). The clinical spectrum of cutaneous infections caused by M. haemophilum appears to be broad (19, 30), varying from localized disease to systemic disease with cutaneous dissemination (49). Multiple skin lesions tend to occur and can present as erythematous papules, plaques, nodules, necrotic abscesses, or chronic ulcers. Cutaneous lesions are found most frequently on the extremities, particularly over joints, and less commonly on the trunk and face. Purpuric and annular lesions have also been described

Several cases of septicemia and pneumonitis due to M. haemophilum have been documented (Tables 2 and 3). The patients with disseminated disease in Table 2 include 11 adults aged 30 to 67 years and 1 6-year-old child. Nine patients were from the United States, one was from Germany, and one was from Brazil. Five patients had AIDS, one had received a renal transplant, one had received a cardiac transplant, two had received a bone marrow transplant, and one was undergoing treatment for multiple myeloma. Only one case of a pediatric disseminated infection has been described (11). A 6-year-old child from The Netherlands with a history of B cell precursor acute lymphoblastic leukemia presented with fever and painful suppurative skin lesions on the knees, elbows, and face. The patient later developed arthritis and osteomyelitis of the right knee in addition to several subcutaneous abscesses, and she remained febrile. Nine patients have been reported to have M. haemophilum pulmonary infections. Six patients were male, and three were

48/M 59/M 52/M 62/M 65/F 17/F 45/F 14 mo/F

67/F

38/F 47/F 37/M 16/M 59/M 27/F 59/F

64/F 42/F

51/M 44/M 59/M 25/F 30/F 51/M 29/F 62/M 51/M 56/M 49/F 53/F 39/M 30/F

35/M

8 22 114 119 72 72 28 28

125

94 94 20 21 149 53 138

138 138

98 85 49 139 108 38 126 47 51 97 97 97 97 80

6

AIDS

Cutaneous vasculitis Sjogren’s syndrome, Crohn’s disease AIDS Renal transplant Diabetes SLE AIDS AIDS AIDS AIDS AIDS Lung transplant Lung transplant Lung transplant Lung transplant AIDS

Autoimmune cirrhosis Myasthenia gravis, corticosteroids AIDS Renal transplant Polymyalgia rheumatica AIDS SLE

IgA deficiency Renal transplant Renal transplant Heart transplant CLL SLE, MDS Renal transplant Unknown immunodeficiency, CD4⫹ ⬍16% RA

Underlying disease(s)

Germany

Italy Taiwan United States Singapore Germany Japan United States Switzerland United States Australia Australia Australia Australia The Netherlands

Singapore Singapore

United States United States Spain United States Germany United States Singapore

Germany

Germany Brazil United States Israel United States United States Venezuela Venezuela

Country

RB, E, CLR; later RB, AZI, L CI, R, CLR CI, RB, CLR CLR, E, I, R Only antiretroviral therapy CLR, E R, CI, CLR, D AZI, RB CLR, TMS, CI I, CLR, P, D, CI CLR, E, R CLR, R, CI, D CLR, R, CI, D (i) D, CI, R, CLR; (ii) Min, RB, E; (iii) I, CI, CY R, E, I, CLR

CLR, CI CLR, D

(i) CLR, CI; (ii) RB, E, CLR; (iii) CLR monotherapy after 8 wk CI, CLR D, RB, AZI I, R, E, AK, CLR, CI, Min E, R, CLR, CI, AK I, E, R RB, CLR, CI (i) CLR, CI, R, I, E; (ii) RB, CLR

CLR, RB, E CLR, CI CI, CLR CI, CLR, R R,CLR, CI R, CLR, G CLR CLR, TMS, R, I

Treatment

Resolution

Resolution Resolution Resolution Resolution Resolution Resolution Died Resolution Resolution Resolution Resolution Resolution Resolution Resolution

Resolved Improvement Resolved Partial resolution Died Improvement Resolved; recurrence and resolution after 2nd course Resolution NA

Resolved

Resolved Resolved Resolved Regression within 3 wk Resolved Resolved Resolved Resolved

Outcome

7 wk, relapse and retreatment with same regimen

5 mo 1 yr NA 6 mo 14 mo 8 mo 11 mo 6 mo NA 42 mo 17 mo 31 mo 18 mo 6 mo

18 mo 18 mo

1 mo 13 mo

6 mo 8 mo 5 mo 14mo

⬎6 mo, NA

6 mo 1 yr 1 yr 1 mo 6 mo NA 6 mo 6 mo

Duration of treatment

LINDEBOOM ET AL.

a I, isoniazid; R, rifampin; RB, rifabutin; E, ethambutol; CY, cycloserine; CI, ciprofloxacin; AK, amikacin; AZI, azithromycin; CLR, clarithromycin; TMS, trimethoprim-sulfamethoxazole; P, pyrazinamide; D, doxycycline; Min, minocycline; G, gatifloxacin; L, levofloxacin; RA, rheumatoid arthritis; MDS, myelodysplastic syndrome; SLE, systemic lupus erythematosus; CLL, chronic lymphocytic leukemia; NA, data not available; M, male; F, female.

Age of patient (yr)/sex

Reference

TABLE 1. Reported cutaneous manifestations in immunocompromised patients, 1996 to 2011a

704 CLIN. MICROBIOL. REV.

VOL. 24, 2011

MYCOBACTERIUM HAEMOPHILUM INFECTIONS

705

76, 135, 156 76, 79 135, 163 117

Reference(s)

136 11

39 121 126 19 123

75, 75, 75, 75, 23

Germany The Netherlands

Country

Cardiac transplant Renal transplant BMT, MDS Multiple myeloma, RA AIDS

Ophthalmologic Manifestations

Subcutaneous nodules Skin lesions, septic arthritis Skin lesions, septic arthritis Subcutaneous nodules NA

Skin, pulmonary, joints NA Pulmonary infiltrate Skin disease Synovial involvement

Skin, pulmonary Skin, pulmonary, joints

Initial presentation(s)

Skin, blood Synovial fluid, blood Skin, blood Skin, blood Blood

Skin, synovial fluid Blood Blood Skin, blood Synovial fluid, blood

Blood, sputum Skin, bone marrow

Culture source(s)

TABLE 2. Reported disseminated infections in immunocompromised patientsa

Age of patient (yr)/sex

States States States

BMT, APML AIDS AIDS AIDS NA

Underlying disease(s)

64/M 6/F

United Brazil United United United

States States States States States

AIDS ALL

46/F 67/M 32/M 51/F 33/M

United United United United United

States

30/F 36/M 37/M 34/M NA

Two reports in the literature described primary ophthalmologic infections due to M. haemophilum (102, 104). Millar et al. (102) described a 55-year-old man with a history of acute myeloid leukemia with chronic bilateral conjunctivitis and dry eyes for a period of 6 months. Skin lesions were also noted on the patient’s face and arms. The clinical condition improved with moxifloxacin and clarithromycin with the addition of valacyclovir and clindamycin 1 week later. Modi et al. (104) presented a unilateral chronic granulomatous iridocyclitis in a 66-year-old man with previous cardiac transplantation and cyclosporine and mycophenolate mofetil treatment. Antibiotic therapy did not prevent the progression of intraocular inflammation, and the patient developed a corneal ulcer that perforated. Enucleation was performed 1 year after the initial presentation. However, the skin lesions regressed with antibiotic therapy. Osteomyelitis A less common manifestation of M. haemophilum in immunocompromised patients is septic arthritis or osteomyelitis with or without cutaneous lesions. Osteomyelitis caused by M. haemophilum resembles that caused by other microorganisms (107, 163). On radiographs, bony resorption with clear margins, cortical destruction, and adjacent soft tissue swelling are apparent. Magnetic resonance imaging (MRI) can reveal wellcircumscribed medullary lesions with cortical disruption and a large soft tissue component (83). Table 4 provides an overview of the cases of skeletal M. haemophilum infections described in the literature. Twenty-six cases have been reported, with a median age of the patients of 45.5 years (range, 20 to 77 years). The underlying illnesses most frequently included AIDS (15 cases) and organ transplantation (7 cases). Uncommon Clinical Presentations

Treatment

RB, E, CLR ⫹ TMS E, R, CLR, AK, drainage of abscesses Imi, CI, CLR, D NA None R, E, CLR, CI E, CLR, CI, AK

CI, CLR, D, I, R, E PAS, R AK, CI, I, CL, D, E, I, R E, I, R NA

Outcome

Cure Cure

Cure NA Died Cure Relapse, died of AIDS Cure Cure Cure Persisted NA

a I, isoniazid; Imi, imipenem; R, rifampin; RB, rifabutin; E, ethambutol; CI, ciprofloxacin; AK, amikacin; CLR, clarithromycin; TMS, trimethoprim-sulfamethoxazole; D, doxycycline; PAS, p-aminosalicylic acid; RA, rheumatoid arthritis; BMT, allogeneic bone marrow transplantation; MDS, myelodysplastic syndrome; APML, acute promyelocytic leukemia; CML, chronic myelogenous leukemia; ALL, acute lymphocytic leukemia; NA, data not available.

female, with a median age of 38 years (range, 27 to 72 years) (Table 3). Eight reports were from the United States, and the most recent report was from The Netherlands. Despite several multidrug regimens (Table 3), a resolution of the infection was observed for less than half of the patients.

Central catheter infections. Ward et al. (152) described an M. haemophilum infection of the central venous catheter tunnel in two young (26 and 29 years old) immunosuppressed patients with hematological malignancy undergoing high-dose chemotherapy supported by bone marrow transplantation. The M. haemophilum infections occurred at the site of the tunneled catheter after the line had been removed. For one patient, therapy consisted of amikacin, clarithromycin, ciprofloxacin, and meropenem for 3 weeks, after which amikacin and meropenem were ceased and ethambutol was started. For the other patient, repeated surgical excisions were combined with clarithromycin, amikacin, and meropenem treatment. The drugs were later changed to rifampin, ciprofloxacin, and clarithromycin. For both patients, the wound eventually healed.

35/M 27/M 27, 75, 76, 135 27, 75, 76, 135, 156

a I, isoniazid; R, rifampin; RB, rifabutin; E, ethambutol; CI, ciprofloxacin; AK, amikacin; AZI, azithromycin; CLR, clarithromycin; P, pyrazinamide; D, doxycycline; Min, minocycline; Ery, erythromycin; S, streptomycin; RA, rheumatoid arthritis; BMT, allogeneic bone marrow transplantation; MDS, myelodysplastic syndrome; AA, aplastic anemia; CML, chronic myelogenous leukemia; BAL, bronchoalveolar lavage; OSAS, obstructive sleep apnea syndrome; NA, data not available.

R, E, CI, AK, D, Ery R, E, AK, P, I, S Skin, sputum Sputum, BAL fluid, lung biopsy specimen Skin, pulmonary infiltrate Pulmonary nodules AIDS BMT, AA

AIDS BMT, MDS AIDS United States United States United States 51/M 42/M 37/M 135 76 76

United States United States

CI, D, E, I, P, R R, E, CI, CLR, AK, P AK, CI, D, E, I, R Sputum, bone BAL fluid, sputum Skin, sputum

Resolution Death Responded and then relapsed and died Death Death

R, AZI None R, E E, I, R, subsequent Min 72/F 32/M 62/F 30/F 137 126 157 123

The Netherlands United States United States United States

RA/OSAS BMT, MDS NA Renal transplant, subsequent AIDS

Pneumonia Pulmonary infiltrate Pulmonary nodule Skin lesions, septic arthritis, subsequent pulmonary involvement Bronchitis Pulmonary infiltrate Skin lesions, pneumonia

Sputum Blood Lung biopsy specimen NA

Resolution Death Cure Initial resolution, subsequent death

CLIN. MICROBIOL. REV.

Reference(s)

Age of patient (yr)/sex

Country

Underlying disease(s)

Initial presentation(s)

Culture source(s)

Treatment

Outcome

LINDEBOOM ET AL.

TABLE 3. Reported pulmonary manifestations in immunocompromised patientsa

706

Epididymal abscess. Keller et al. (73) described an epididymal abscess due to M. haemophilum infection in a renal transplant patient. A right orchidectomy was performed, combined with clarithromycin, rifabutin, and ethambutol treatment. The symptoms resolved over 5 months of follow-up. Mixed infections. Dual infections with M. haemophilum and other NTM species are extremely rare but also difficult to diagnose. Since the first case report by Branger et al. (12), describing a mixed infection with M. haemophilum and M. xenopi, two new cases have been reported. Bekou and colleagues (8) reported a skin infection with multifocal nodules of variable sizes arranged in a sporotrichoid-like manner on the extremities and back of a 48-year-old male patient with an IgA deficiency. Both M. haemophilum and M. kansasii were cultured. Treatment with clarithromycin, rifabutin, and ethambutol for 6 months led to a complete clinical remission of the skin lesions. Phowthongkum et al. (111) described a 40-year-old male patient with AIDS who developed a spindle cell pseudobrain tumor as a result of M. haemophilum and M. simiae infection. He was treated with isoniazid, rifampin, pyrazinamide, ethambutol, and clarithromycin. One month after hospitalization, he commenced antiretroviral treatment, including zidovudine, lamivudine, and efavirenz. He was discharged home, and was seen for the last time 3 months after the operation.

IMMUNOCOMPETENT PATIENTS Adult Infections Cervicofacial infections. An outbreak of 12 cases of M. haemophilum skin infection with lymphadenitis after permanent makeup on the eyebrows was described recently (52). The ink used by the tattoo artist was found to be contaminated with M. haemophilum. All 12 patients were female, with a median age of 56 years, and none of the patients were immunosuppressed. The patients presented with an inflammatory lesion consisting of a few red papules or pustules or an erythematous plaque on one eyebrow. In all cases, the lesion was associated with ipsilateral lymphadenopathy in the parotid region, affecting one or more lymph nodes (median, 2; range, 1 to 5). Eight patients presented with an abscess, which later developed into a fistula in seven cases, whereas none of the patients reported systemic symptoms. Minani et al. (103) described a 27-year-old immunocompetent woman with a right buccal abscess and submandibular lymphadenitis. The patient was cured with surgical excisional therapy of the affected lymph nodes and drainage of the buccal abscess. A retrospective overview of another six patients with cervicofacial lymphadenitis (five females and one male, with an age range of 19 to 65 years) seen over a 15-year period in Phoenix, AZ, was also presented (103). “Other” skin lesions. Skin lesions in immunocompetent adults due to M. haemophilum infection are rare and the result of injury. Two cases have been reported (99, 128): a 61-yearold male who sustained several lacerations to the forearm when he was thrown against coral while surfing and a 65-yearold female who developed subcutaneous skin nodules after coronary artery bypass surgery.

707 MYCOBACTERIUM HAEMOPHILUM INFECTIONS VOL. 24, 2011

Lymphoma Renal transplant Cardiac transplant AIDS AIDS France

Australia Australia South Africa United States United States

OMb

Partial response Died Died No improvement No improvement

Outcome I, R, I, R, NA R, I, I, R,

Cure

Therapy

Surgery; Min, Ery for 2 mo; I, R, E, Min for 6 mo R, I, P, E

Other site(s)

Skin, blood

R, Min I, R, E, CI, CL, AK

Improvement, stable at 19-mo follow-up Resolution Initial improvement, relapse in 6 wk Resolution

E E

Skin Skin, sputum

R, I, E, AK, CI, CL, P

Resolution No improvement

AIDS

United States

United States

Knee

Hand

Skin

Skin, pulmonary

E, CLR, CI, AK

I, R, E, Min

P, E, ET E

Skin

R, I, P, E for 14 mo I, R, E

Improvement in 10 wk Resolved 9 mo later No improvement

NA Blood Soft tissue abscess, BAL fluid Skin

Skin Skin, lungs

CL, AK, D, R R, I, P R, I, P, E, A, CI, CL

46/M Renal transplant/AIDS

United States

Hand

R 3rd finger, R calcaneus L ankle Finger, toes, tibia, elbow, T9-10 vertebrae L foot R olecranon R ankle and tibia

L ankle, tibia Bilateral tibia and fibula

L middle finger, L knee

Foot Ankle Limbs

TABLE 4. Reported septic arthritis/osteomyelitis in immunocompromised patientsa

58/M 55/F NA 32/M 34/M Renal transplant United States

Area(s) of septic arthritisb

101 101 26 96 117 48/M AIDS United States United States L knee

Country

54 36/M AIDS AIDS United States

Bilat knees

Underlying disease

79 21/F 44/M AIDS United States France

R elbow Knees, ankles

Age of patient (yr)/sex

57 33 31/F AIDS AIDS Australia United States United States

Reference(s)

163 37/M NS AIDS AIDS AIDS

Ankles, L wrist R finger

76, 135, 163 118 39/M 41/M 49/M

65 30/F AIDS

United States Hip

R knee

133 132 64

CI, RB, CY, AZI

123 33/M T cell lymphoma

United States

Ankle

Skin Skin, blood, lymph nodes

123 77/M

RA (corticosteroids)

United States

Foot

123 66/F AIDS

Finger

Skin, pneumonia

E, R, CLR CI, RB, CLR

CI, RB CLR, R NA CI, CLR, D, R (⬎6 mo) Imi, CI, CLR, D, 2 mo

Cure Cure

Died 2 mo after initial presentation Cure Cure NA Improved Resolution NA

Skin

D, R, subsequent excision ⫹ D, R NA

Curettage

123 56/M

States States

Hand

Improved after treatment Resolution of lesion, died Died of AIDS complications Relapse after 1 yr, died of lymphoma complications Cure

123

United United Brazil United United

Tibia R wrist, R ankle

Skin

Renal transplant Cardiac transplant AIDS AA/BMT Cardiac transplant

Germany Canada

Olecranon Elbow NS

45/F 20/M 30/M 47/F 46/F

AIDS Polycythemia vera

Wrist, knees, ankles

123 112 121 126 39 53/F 56/F

States States

56 37

a I, isoniazid; Imi, imipenem; R, rifampin; RB, rifabutin; E, ethambutol; ET, ethionamide; CY, cycloserine; CL, clofazimine; CI, ciprofloxacin; AK, amikacin; AZI, azithromycin; CLR, clarithromycin; P, pyrazinamide; D, doxycycline; Min, minocycline; Ery, erythromycin; RA, rheumatoid arthritis; AA, aplastic anemia; BMT, bone marrow transplant; OM, osteomyelitis; NA, data not available. b L, left; R, right.

708

LINDEBOOM ET AL.

FIG. 2. Clinical picture of a child with a cervicofacial Mycobacterium haemophilum lymphadenitis presenting as a fluctuant swelling with red skin discoloration.

Pediatric M. haemophilum Infections Cervicofacial infections. Lymphadenitis is the most common clinical manifestation of NTM infection of children (155). Since the first reported case of cervicofacial lymphadenitis in an immunocompetent child in 1981 (31), seven additional cases of children with head and neck lymphadenitis have been added to the literature (3, 123, 141, 147). M. haemophilum was recently reported to be a major cause of lymphadenitis in immunocompetent children in Israel and The Netherlands (90, 164). These reports showed that M. haemophilum is the second most commonly recognized pathogen in children with cervicofacial NTM lymphadenitis. Patients with M. haemophilum lymphadenitis tended to be older than patients with the more common M. avium lymphadenitis (25, 90). In the study from The Netherlands (90), the M. avium-infected and M. haemophilum-infected patients did not differ with respect to sex, duration of lymph node swelling prior to presentation, or clinical symptoms, but M. haemophilum infections of the head and neck were associated with an infection of multiple lymph nodes (Fig. 2, 3, and 4) and the involvement of extranodal areas, such as the medial canthus, cheek, or ear lobe (60, 90, 92). Children with M. avium or M. haemophilum cervicofacial lymphadenitis seldom exhibited general clinical symptoms (90, 164), although some children experienced a loss of appetite. As a result of a diagnostic delay, most children with M. haemophilum lymphadenitis (80%) presented in a secondary or tertiary center in the stage of lymph node fluctuation with discoloration of the skin. Inguinal lymphadenitis. One case of a 5-year-old girl with a painful, enlarged lymph node in the groin has been reported (89). The portal of entry was most likely a wound on the dorsum of her foot. During antimycobacterial therapy with clarithromycin and rifabutin, the inguinal lymph node started suppurating, and after 12 weeks of treatment, complete necrosis of the lymph node was visible. The surgical excision of the affected inguinal lymph nodes led to complete resolution. Pulmonary involvement. Armstrong et al. (3) described a 12-month-old male infant with a 6-week history of daily fever, anorexia, and weight loss. Examination revealed fever, cough,

CLIN. MICROBIOL. REV.

FIG. 3. Clinical picture of Mycobacterium haemophilum lymphadenitis after skin breakdown.

tachypnea, tachycardia, and decreased breath sounds over the right upper lobe of the lung. No immunodeficiencies were detected, and after mediastinal biopsy, antituberculous medication with pyrazinamide, rifampin, isoniazid, and pyridoxine reduced the clinical symptoms. After 6 weeks, the antibiotic therapy was changed to erythromycin, which was prolonged for 15 months, with a final resolution of the disease. ANIMAL INFECTIONS M. haemophilum infection is not restricted to a human host. M. haemophilum appears to be pathogenic in fish and has caused clinical manifestations in a snake and a bison similar to those seen in humans (63, 69, 74, 154). A royal python was diagnosed with pulmonary mycobacteriosis caused by both M. marinum and M. haemophilum (63). Normal lung tissue was largely replaced by granulomatous tissue containing necrotic foci, as is often observed for mycobacterial disease in humans. Cultures of tissue biopsy specimens contained numerous AFB

FIG. 4. Ulcerating open wound as a result of a cervicofacial Mycobacterium haemophilum infection.

VOL. 24, 2011

MYCOBACTERIUM HAEMOPHILUM INFECTIONS

representing both species. Another report described an intradural mass compressing the spinal cord in a bison (69). Again, histological examination showed necrotic granulomatous tissue containing a large number of AFB. 16S rRNA gene sequencing analysis of the mycobacterial culture identified M. haemophilum. M. haemophilum appears to be highly pathogenic in zebrafish, as several outbreaks have been reported (74, 154). At least three unrelated outbreaks, with mortality rates of up to 20%, were caused by this species. All organs seemed to be infected, and massive amounts of bacilli were observed in granulomas and throughout regions of diffuse inflammation. PATHOGENESIS M. haemophilum infections are similar to those caused by M. marinum and M. ulcerans; they occur most commonly as necrotic lesions within the regions of the body with the lowest temperatures (19). Histological examination usually reveals a granulomatous reaction with necrotic foci. M. haemophilum is apparently of low virulence, as most healthy mice and guinea pigs in earlier studies survived for an observation period of 3 months after intramuscular, intravenous, and subcutaneous inoculations of large numbers of bacilli (130, 131). However, some of the mice died after 2 to 4 weeks, with large numbers of AFB in liver, spleen, and kidneys. The intramuscular injection of M. haemophilum into the thighs of frogs did not result in abnormalities when the frogs were kept at room temperature. However, the animals died within 20 days when kept at 30°C, with M. haemophilum infestation in the liver and kidneys. In vitro, M. haemophilum seems to have a preference for growth in cultured human endometrial carcinoma cells (Hec-1-B), compared to human microvascular endothelial cells (HMEC-1) (43, 44). An epithelial cell culture infection model suggested greater intracellular replication at 33°C than at 37°C and showed that the bacilli are associated with cytotoxicity at the lower temperature (43, 44). These observations indicate that M. haemophilum is a facultative intracellular bacterium. Additionally, M. haemophilum exhibits contact-dependent cytolytic activity at 33°C, similar to the effect observed for M. tuberculosis infections. Thus, the pathogenicity of M. haemophilum appears to be temperature dependent, which is consistent with infection and tissue damage in skin and other superficial body sites with a lower temperature. EPIDEMIOLOGY Typing of M. haemophilum Several Mycobacterium species have been examined extensively by molecular typing, but limited information is available on the genetic diversity of M. haemophilum. Three typing studies have been conducted to date, based on pulsed-field gel electrophoresis (PFGE) (162), restriction fragment length polymorphism (RFLP) analysis (77), and amplified fragment length polymorphism (AFLP) analysis (16). All three methods demonstrated a high degree of clustering among the clinical isolates investigated, and a sufficient degree of discrimination was observed among isolates that were not epidemiologically related. PFGE and RFLP analysis were used

709

to type isolates from the United States, most of which came from the New York City area. In the AFLP study (16), isolates from different continents were tested, including the strains from the United States that were also subjected to RFLP analysis and PFGE. The general conclusion from these three studies was that a high degree of clustering exists among isolates from the same geographic area and that a high degree of genetic stability is present over time. Clusters of identical DNA fingerprint types were observed within close geographical proximity, but the isolates were not necessarily derived from the same hospitals and not found in geographically distant locations. Genetic conservation was also demonstrated by several clusters of clonal types for extended time periods; one cluster from New York linked isolates over a period of 16 years, and two clusters from Australia remained unchanged for 15 and 18 years, which suggests an extremely low evolutionary rate for this mycobacterium. This bacterium may survive in a highly suitable niche, such as tap water, without any selective pressure. Although the typing results of the three studies are in accordance and technically reliable, typing results should be analyzed with caution because isolates with (nearly) the same DNA fingerprinting profiles are not necessarily epidemiologically linked. Whole-genome sequencing of multiple strains will facilitate the establishment of a robust and detailed phylogenetic tree that may serve to clarify the epidemiology of M. haemophilum infections in humans and the environment. This method was recently shown to be highly informative when it was applied to an M. tuberculosis outbreak in which two separate lineages were identified to occur simultaneously in one social network (48). Environmental Findings Although no clinical isolates have been linked directly to environmental isolates, several findings suggest that water reservoirs are a likely source of M. haemophilum infection. For a cluster of M. haemophilum infections in New York, the hospital drinking water supply was suspected to be the common source, but this was not proven (T. E. Kiehn, Memorial Sloan Kettering Cancer Center, New York, NY, personal communication). The resistance to common disinfectants, temperature tolerance, and ability to form biofilms exhibited by mycobacteria are all preferential characteristics for survival and persistence in water systems and reservoirs (41). One paper describing an M. haemophilum infection in a patient after sustaining a coral injury suggested that seawater or coral is also an environmental source (128). Several studies have been conducted with the objective of investigating the presence of NTM in water systems (146) However, the specific requirements for the detection of M. haemophilum were often not met in these studies. For example, Covert et al. (27) employed molecular identification after culturing without specific requirements for M. haemophilum. Chang and colleagues (24), using a PCR-RFLP method for the direct detection of mycobacteria in water samples, showed a high prevalence of AFB. However, the reverse primer sequence used in that study did not match the M. haemophilum sequence, and thus, direct detection was compromised. Molecular detection using concentrated water samples containing

710

LINDEBOOM ET AL.

AFB was unsuccessful overall, and the method was eventually applied to the identification of isolates cultured without the culturing requirements necessary for M. haemophilum. Both studies showed that a variety of mycobacterial species were present in chlorine-treated water supplies and were thorough, but M. haemophilum might have been overlooked. Only a few studies allowed the detection of M. haemophilum by molecular methods or specific culturing methods (40, 68, 113, 154). Hussein and colleagues (68) did include species detection, but they encountered only other NTM. Three studies detected M. haemophilum. Falkinham et al. (40) found it in three samples, comprising one water sample and two biofilm samples, all from different water distribution systems in the United States. Whipps et al. (154) detected M. haemophilum in biofilms from four zebrafish tank meniscuses and one tank drain, all from a zebrafish research center in which M. haemophilum caused significant mortality among the fish population. Pryor et al. (113) cultured M. haemophilum from a water distribution system (unknown sample type) as one of many other Mycobacterium species. In one publication, an environmental M. haemophilum isolate not directly associated with water was described. Mycobacterial isolates were cultured from the intestines and surface of hospital cockroaches in Taiwan, and M. haemophilum was found on the surface of one cockroach (109). DIAGNOSTICS Skin Testing No specific antigen test is available for M. haemophilum infections, although in the past, purified protein derivatives (PPDs) of M. avium, M. kansasii, M. scrofulaceum and M. marinum, M. intracellulare, M. gordonae, and M. fortuitum have been used for the diagnosis of NTM infections. Unfortunately, a few years ago the production of NTM-PPD (Statens Serum Institute, Denmark) was terminated, although skin testing appeared to be useful for the diagnosis of NTM infections in children. Because of cross-reactivity between the immune reactions to PPDs of different species, the tuberculin-PPD test often shows false-positive reactions due to previous encounters with NTM (91). The problem with previous NTM encounters is not expected in young children; therefore, a positive tuberculin test can be indicative of NTM disease in this patient group, except for children living in a country where tuberculosis is highly endemic. For the initial diagnosis of NTM lymphadenitis, the tuberculin test has an optimal cutoff value of 5 mm for a positive skin induration (91). Using a 5-mm cutoff, the tuberculin PPD has 71% sensitivity for M. haemophilum and a 98% positive predictive value (PPV). Using a 10-mm cutoff (the induration cutoff considered positive for M. tuberculosis reactivity), 57% of all confirmed M. haemophilum infections yielded positive skin indurations. Histopathology Tissues infected with M. haemophilum show, almost without exception, granulomatous infiltrates with necrosis (19, 35). The granulomas comprise variable forms of granulocytes, lymphocytes, monocytes, and multinucleated giant cells. Bacilli can be

CLIN. MICROBIOL. REV.

observed both extracellularly and intracellularly, and they can be abundant or scarce in affected tissue (19, 35, 126). No specific clinical and histological manifestations can be attributed to M. haemophilum. M. haemophilum skin infection often mimics M. marinum infection: it forms erythematous papules or nodules, often overlying or above the joints, and in later stages, it becomes suppurative/ulcerative. However, in contrast to M. marinum infections, the nodules are painful, and sporotrichoid spread is seldom seen in M. haemophilum infections (19). Skin manifestations sporadically include lichenoid dermatitis, panniculitis, vasculitis, or annular plaques. Histological findings for 16 skin biopsy specimens from 11 immunocompromised patients with culture-proven M. haemophilum infections revealed most commonly (7 of 16 biopsy specimens) a mixed histopathological pattern of suppurative and granulomatous reactions (19). Four biopsy specimens showed well-formed epithelioid granulomas. The authors of that study noted that infections by M. haemophilum can also present with nongranulomatous or paucigranulomatous reactions without necrosis, probably due to the immunocompromised state of the patients. Microscopy M. haemophilum is a strongly acid-fast bacterium and can be stained with Ziehl-Neelsen, modified Kinyoun, or auramine dye. The bacilli appear as short, and often curved, rods (1.2 ␮m to 2.5 ␮m in length) and can be pleomorphic. No specific growth or morphological differences exist between this and other species. Because M. haemophilum has the tendency to clump, a stain from a cultured isolate can exhibit strings of AFB, as is sometimes attributed exclusively to M. tuberculosis. Cord formation or cording should no longer be attributed exclusively to isolates of M. tuberculosis, as has recently also been demonstrated for nonpathogenic mycobacteria (71). Culture Like most of the pathogenic Mycobacterium species, M. haemophilum is slowly growing. Visible growth can take as long as 8 weeks. The normal growth temperature for mycobacteria is 35°C to 37°C. M. haemophilum, however, prefers a lower growth temperature of 30°C to 32°C and requires iron supplements such as hemin or ferric ammonium citrate, which can be added to both liquid and solid media (32, 122). Culturing of mycobacteria is most frequently applied to a system measuring the assimilation of bacteria in broth medium such as the BBL Mycobacteria Growth Indicator Tube (MGIT) containing Middlebrook 7H9 medium. A combination of a liquid culture medium with a solid medium is recommended. Solid egg-based media such as Lo ¨ wensteinJensen (LJ), Coletsos, Stonebrink, Herrold’s, or Dubos medium and solid agar-based media such as Middlebrook 7H10 and 7H11 agars are commercially available but must be supplemented with iron or hemin to allow the growth of M. haemophilum, as previously described (4). Growth enhancers, such as mycobactin and OADC (containing oleic acid, albumin, dextrose, catalase, and NaCl), and antibiotics to inhibit the growth of contaminants are often

VOL. 24, 2011

MYCOBACTERIUM HAEMOPHILUM INFECTIONS

added: PANTA (containing polymyxin, amphotericin B, nalidixic acid, trimethoprim, and azlocillin) and/or PACT (containing polymyxin B, amphotericin B, carbenicillin, and trimethoprim) (129, 158). The effect of these growth enhancers or antibiotic supplements on M. haemophilum has not been examined. The application of a decontamination protocol prior to culture helps to further decrease contamination with commensals and to release culturable bacilli from tissue (17). Several decontamination protocols are available, but it should be considered that most of them also decrease to some extent the recovery of mycobacteria. In our institute, we follow a NALC (N-acetyl-L-cysteine)-NaOH procedure for those samples that are contaminated and culture positive for rapidly growing bacteria on a standard blood agar medium (13).

Molecular Identification Methods M. haemophilum can easily be differentiated from other species by sequencing. The representation of the species in the publicly available GenBank databases is sufficient for identification. Complete or partial ITSs and 16S rRNA, rpoB, and hsp65 genes represent 28 of 48 M. haemophilum sequences submitted to the database to date (January 2011). For most other housekeeping genes, only one sequence is available. Although the genetic marker most suitable for species identification is still unclear, all sequence targets in the database enable the identification of M. haemophilum. A few commercial assays are available for the identification of cultured NTM isolates. Two reverse line probe assays include M. haemophilum: the GenoType Mycobacterium AS (Hain Lifescience GmbH, Nehren, Germany) (115) and the Inno-LiPA-Mycobacteria V2 (Innogenetics, Ghent, Belgium) (143, 144) assays. Other assays do not include the species, such as the AccuProbe assay, a chemiluminescence assay (Gen-Probe Inc./bioMe´rieux, Marcy l’Etoile, France), and the Speed-Oligo Mycobacteria assay, a hybridization dipstick test (Vircell, Spain). The newest software and database versions of Microseq 500 ID (Microseq ID 16S rDNA Full Gene Library v2.0, Applied Biosystems, Foster City, CA), a sequencing system, include a database with 86 mycobacterial species, including M. haemophilum (Applied Biosystems). Also, several noncommercial molecular assays have been developed to differentiate between Mycobacterium species and include M. haemophilum. High-performance liquid chromatography (HPLC) has also been successfully applied (140) The new assays either employ species-specific probe hybridization, such as array probes (142, 161), or use restriction patterns to differentiate between species (118). Newly developed methods that are currently being evaluated for application as tools to identify bacterial isolates might be applicable for the identification of species of Mycobacterium isolates. Matrix-assisted laser desorption ionization–time of flight mass spectrometry (MALDI-TOF MS) and Raman spectrometry (18, 95, 120) as well as the new-generation sequencing method pyrosequencing (145) have been described for the differentiation of NTM species. Although M. haemophilum has been included in the NIH database, clinical isolates have not yet been tested (120).

711

Direct Detection Methods The direct detection of NTM as a group is still being challenged, as only a few molecular assays have been described and validated for direct application to clinical materials (5, 13, 67, 116, 134). Only two of these assays have been applied to the detection of M. haemophilum in clinical materials. Conventional PCR and subsequent restriction analysis (PRA) of hsp65 in all Mycobacterium species were applied successfully to biopsy specimens from four patients with M. haemophilum skin infections (28, 151). A 439-bp fragment was amplified and digested into species-specific band patterns by two restriction enzymes. However, the assay includes the handling of PCR products and therefore poses a contamination risk. The second assay is a real-time PCR assay targeting the ITS between the 16S rRNA and 23S genes of all slowly growing Mycobacterium species (13). Mismatches in the forward primer and genus-specific probe have been encountered in several rapid-growing mycobacteria; therefore, for the detection of this group of species, this assay is less proficient. A speciesspecific probe subsequently enables the recognition of M. haemophilum. M. haemophilum-specific culture was found to be less sensitive than the real-time PCR assay when applied directly to biopsy specimens from children with cervicofacial lymphadenitis (14). Of 16 patients with evidence of M. haemophilum infection, 9 (56%) were positive by auramine staining, and 9 (56%) were positive by M. haemophilum-specific cultures. Thirteen specimens (81%) were positive by genus-specific detection, 11 of which were also positive by M. haemophilum-specific detection. This assay was also applied to formalin-fixed/paraffin-embedded biopsy specimens from patients with granulomatous inflammation of the skin, which were stored between 1984 and 2004 (15). Of 30 patient materials tested, 13 (43%) were found to contain mycobacterial DNA. Only 5 of the patients had been previously diagnosed with a mycobacterial disease. M. haemophilum was identified as the most common species (n ⫽ 7). In this study, PCR was not compared with conventional techniques. Another possible approach for direct detection is the application of generic PCR targeting a Mycobacterium-specific fragment that is subsequently sequenced to identify the involved species. This approach was applied in several reported M. haemophilum cases (52, 70, 114). The method can be performed by using a number of gene fragments (see “Molecular Identification Methods” above). Diagnostic Approach M. haemophilum infection should be considered for immunocompetent patients with nonpyogenic cervicofacial lymphadenitis. M. haemophilum can induce reactions in the tuberculin PPD skin test similar to those induced by M. tuberculosis and could be misdiagnosed when positive culture results are lacking (3, 59, 91). In general, a 10-mm tuberculin PPD cutoff point is recommended for the identification of latent M. tuberculosis infections, whereas a reaction of 5 to 9 mm is more likely to indicate NTM infection (45, 46). Therefore, although it is not decisive, the tuberculin PPD test can be helpful as a

712

LINDEBOOM ET AL.

diagnostic tool with an induration cutoff of ⬎5 mm as an indication of NTM infection in children. M. haemophilum involvement should also be suspected for immunocompromised patients with typical NTM manifestations combined with skin lesions. Specific M. haemophilum detection should be carried out concurrently with standard mycobacterial detection for clinical samples obtained from superficial body sites, such as skin biopsy specimens and superficial lymph node biopsy specimens. Overall, the failure to isolate a pathogen from clinical specimens with positive acid-fast stains should prompt a targeted search for M. haemophilum using appropriate culture conditions and molecular techniques. A full diagnostic regimen for the optimal detection of M. haemophilum in biopsy specimens includes acid-fast staining, mycobacterial culturing at two temperatures using media with and without iron additives, and molecular detection. The diagnosis of mycobacterial infection by the direct detection of the pathogen is achieved by use of fine-needle aspiration biopsy (13), excision of the affected tissue, or respiratory specimens. After decontamination using, for example, the NALCNaOH decontamination protocol, biopsy specimens should be stained with auramine and investigated microscopically, followed by standard mycobacterial culturing at 35°C in liquid MGIT medium and on solid LJ medium. In addition to this generic protocol, M. haemophilum-specific culturing should be performed at 30°C on LJ medium supplemented with iron citrate (preferably combined with a liquid medium using hemin supplementation). Because culture for M. haemophilum is less sensitive than the real-time PCR assay described above (14), molecular diagnosis should also be attempted, preferably using genus-specific detection and M. haemophilum-specific detection. Molecular detection also enables biopsy specimens and other histopathological materials to be examined for the presence of mycobacterial DNA when culturing is not possible due to tissue fixation (15). This approach offers an excellent opportunity to investigate the presence of newly identified Mycobacterium species in stored patient materials. However, positive PCR results need to be interpreted with caution. The widespread presence of NTM in the environment may result in the contamination of patient samples with bacilli or DNA fragments during processing. Thus, the application of a highly sensitive NTM DNA detection method can result in false-positive results. ANTIMICROBIAL SUSCEPTIBILITY No standardized procedure is available for the susceptibility testing of M. haemophilum, although a recent CLSI document includes recommendations for a disk agar elution method for M. haemophilum (24a). The application of different culture media can result in variations in the MIC values obtained for the same isolate. Moreover, European and U.S. guidelines do not always fully agree on the critical concentrations and protocols for susceptibility testing (160). Therefore, the in vitro susceptibilities presented in Table 5 are approximations. M. haemophilum appears to be susceptible to ciprofloxacin, clarithromycin, rifabutin, and clofazimine but resistant to isoniazid and ethambutol (96, 105, 126, 141). Discrepant results have been observed for amikacin

CLIN. MICROBIOL. REV. TABLE 5. Resistance of clinical isolates to antimicrobial agentsd

Antimicrobial agent

1993 study (n ⫽ 12)a

MIC50

MIC90

Ciprofloxacin 2 8 Clarithromycin ⱕ0.25 ⱕ0.25 Rifabutin ⱕ0.03 ⱕ0.03 Rifampin 0.5 1 Amikacin 4 8 Ethionamide R* R* Streptomycin NT NT Ethambutol R* R* Isoniazid 8 ⬎32 Clofazimine 2 2 Prothionamide NT NT Cycloserine NT NT

2001 study (n ⫽ 16)b Disk (␮g/ml)

% sensitivity

2 3 NT 1 2 5 10 5 0.2 NT NT NT

100 100 NT 94 100 0 100 0 0 NT NT NT

CHIMED study, 2003–2004 (n ⫽ 18)c MIC50

MIC90

ⱕ1 4 ⱕ2 ⱕ2 ⱕ0.2 ⱕ0.2 0.2 1 10 20 NT NT 10 20 ⬎20 ⬎20 ⱖ20 ⱖ20 ⱕ0.5 ⱕ0.5 5 20 50 ⬎50

a Data from reference 9. The method applied was a microtiter array with Middlebrook 7H9 broth plus hemin. MICs are in ␮g/ml. b Data from reference 126. The method applied was a disk elution method on Middlebrook 7H10 agar with hemin. c The method applied was an agar dilution method on Middlebrook 7H10 medium with a hemin source. MICs are in ␮g/ml. d R*, tested but not active; NT, not tested.

and streptomycin; our results demonstrate high MIC values, allowing us not to consider aminoglycosides for the treatment of M. haemophilum infections. Isoniazid may be more active than indicated by the in vitro test results, since hemin, used as a broth supplement, can antagonize the in vitro activity of isoniazid (9). Interesting results were obtained for cycloserine, with an MIC50 of 50 ␮g/ml. While macrolides and rifamycin appear to be highly active against M. haemophilum, resistance is readily acquired by a single mutation in the 23S gene and the rpoB gene, respectively (78, 110). Therefore, dual or triple therapy is advised over monotherapy.

TREATMENT No standard guidelines are available for the treatment of M. haemophilum infection. Although no optimal therapeutic regimen and treatment duration for M. haemophilum have been established, experts generally agree that patients should be placed on multiple antibiotics that include some combination of clarithromycin, ciprofloxacin, and one of the rifamycins (123, 126) for a duration of 12 to 24 months (126). Therapy should be tailored to the individual patient based on his or her disease presentation and underlying degree of immune suppression. The contribution of antibiotics to the healing of M. haemophilum lesions is difficult to evaluate. Recovery may depend mostly on an improved immunologic state (108). Even if M. haemophilum infections are diagnosed early, adequate treatment may be complicated by an inability to reduce immune suppression, adverse reactions to the antibiotics, patient intolerance of the antibiotics, the antimicrobial resistance of M. haemophilum isolates, interactions between antimicrobials and immunosuppressive agents, and superinfection of cutaneous lesions with, for example, Staphylococcus aureus (39).

VOL. 24, 2011

MYCOBACTERIUM HAEMOPHILUM INFECTIONS

Immunocompromised Patients Skin lesions. Only a few M. haemophilum infections in immunocompromised children have been described (11, 21, 28). One patient was cured after antibiotics for 6 months (28), whereas the other two patients failed treatment (11, 21). One of these failing patients was cured after immune restoration, surgical drainage, and additional antibiotic treatment (11). Numerous cases of skin infections in immunocompromised adults have been reported (Table 1). In all patients for whom therapy was reported, treatment consisted of antituberculous drugs guided by the susceptibility pattern of the cultured microorganism. The regimen usually consisted of at least three drugs: almost always clarithromycin (29) plus ciprofloxacin, ethambutol, and/or rifabutin-rifampin. The treatment duration varied between 3 and 42 months, with a median of approximately 6 months. For AIDS patients, highly active antiretroviral therapy (HAART) was usually also started. With one exception (47), all patients were cured. To summarize, antibiotic treatment is indicated for patients with M. haemophilum skin infections. Curative surgical excision is possible in rare cases with few infected sites (100). The duration of antibiotic therapy is not well defined and depends on the clinical presentation, degree of immune suppression, and clinical course. In an earlier review the minimum recommended duration of antibiotic therapy was 12 months, but treatment may need to be extended for up to 24 months (123, 126). The exacerbation of the skin lesions shortly after the initiation of treatment, however, is not uncommon. These exacerbations most likely occur as a result of a paradoxical reaction: an immune response to the local release of products of mycobacterial cell death and lysis. These reactions tend to improve within 2 to 3 weeks (85, 104, 126). In general, patient outcomes tend to be satisfactory for M. haemophilum skin infections (126). Disseminated infection/pulmonary infection. Tables 2 and 3 give an overview of the reported cases of disseminated and pulmonary infections and the subsequent treatment. Five out of the 10 patients reported with disseminated disease responded to treatment. For disseminated M. haemophilum infections, a multidrug regimen combining clarithromycin, ciprofloxacin, and rifampin-rifabutin is recommended (157). For the reported pulmonary infections (Table 3), the level of response to treatment is lower. Only three patients from the nine reported cases responded permanently to the therapeutic regimen. Although no studies of the duration of treatment for M. haemophilum infections have been conducted, American Thoracic Society guidelines recommend treatment until cultures taken during therapy are negative for 1 year (55). Whether tumor necrosis factor alpha (TNF-␣) treatment can be continued during antimycobacterial treatment is a matter of debate (148). In active tuberculosis infections, treatment with TNF-␣ is contraindicated until patients complete a standard regimen of antituberculosis therapy. No information is available for NTM disease (148). Pyomyositis. The majority of the described cases of pyomyositis were successfully treated with a combination of surgery and antibiotic therapy. Surgical debridement of necrotic tissue is required when extensive inflammation is present (124). Based on limited data from individual cases (70, 82, 124), a

713

combination of surgery and antibiotic therapy with clarithromycin, ciprofloxacin, and one of the rifamycins appears to be effective. However, the wounds did not resolve in a case reported by Shih et al. (127). The therapeutic regimen consisted of ethambutol, rifampin, clarithromycin, ciprofloxacin, and amikacin. Repeated debridement of the right thigh was performed, but the patient died of fungemia due to Candida glabrata 3 months after admission. The duration of therapy should depend on the patient’s underlying disease presentation, degree of immunosuppression, and response to therapy. Treatment should generally be continued for at least 1 year and perhaps for as long as 2 years (123, 126). Skeletal infections/osteomyelitis. Data from the publications on skeletal M. haemophilum infections are presented in Table 4. The clinical response to treatment varies, even when the above-mentioned antibiotics are used. Prolonged maintenance therapy lasting months, or even years, with several drugs is generally necessary, particularly for patients with sustained immunosuppression (83).

Immunocompetent Patients Immunocompetent adults. In adults, the most frequently reported manifestations are skin lesions with or without lymphadenitis. Success has been reported with antibiotic treatment for 4 to 6 months (8, 124, 128). Treatment in these patients consisted of clarithromycin, rifabutin, and ethambutol or ciprofloxacin. In a recent case series of 12 patients with eyebrow lesions and cervicofacial lymphadenitis, surgical excision was curative, and in the majority of the cases antibiotics were not successful (52). Immunocompetent children. The most common manifestation of M. haemophilum infection in immunocompetent children is cervicofacial lymphadenitis. Excisional surgery leads to a quick resolution and the best esthetic outcome (87, 88). In more advanced stages with extensive necrosis and skin discoloration, excisional surgery can be technically difficult. Whether antibiotic treatment offers benefits over observation alone in these cases is not clear. Both success and failure have been reported for antibiotic treatment (60, 88, 92, 147). In the largest reported case series, 32 children in Israel were treated by observation alone (164). Total resolution was achieved for 71% of patients within 6 months and for the remaining patients within 9 to 12 months. For children with an advanced stage of nontuberculous mycobacterial cervicofacial lymphadenitis, no significant difference in the median healing times between an observational approach and antibiotic therapy with clarithromycin and rifabutin was found (86).

Treatment Outcome In the cases reported after 1996, almost all immunocompromised patients were cured, with the few exceptions described above (47, 104, 126, 127, 149). Thus, in summary, most immunocompromised patients will recover after prolonged antimycobacterial treatment, especially those with skin infections, but mortality can occur in patients with deep-seated infections. In these circumstances, surgery is usually not an option, leaving

714

LINDEBOOM ET AL.

only an alleviation of immunosuppressive medication, if possible, or, in AIDS patients, antiretroviral treatment. RECOMMENDATIONS AND CONCLUSION In conclusion, M. haemophilum infection should be considered in the differential diagnosis of chronic cervicofacial lymphadenitis in young immunocompetent children and ulcerating skin lesions and/or arthritis in immunocompromised patients, especially when AFB are seen by direct microscopy and when routine mycobacterial cultures remain sterile. Detailed clinical information, adjusted mycobacterial culture procedures, and molecular techniques all contribute to the adequate and rapid diagnosis of M. haemophilum infections. The outcome of M. haemophilum lymphadenitis in immunocompetent patients favors surgical intervention rather than antibiotic treatment. REFERENCES 1. Abbott, M. R., and D. D. Smith. 1981. Mycobacterial infections in immunosuppressed patients. Med. J. Aust. 1:351–353. 2. Abell, F., P. B. Harrison, and M. Seldon. 1994. Mycobacterium haemophilum infection in an elderly patient. Aust. N. Z. J. Med. 24:404. 3. Armstrong, K. L., R. W. James, D. J. Dawson, P. W. Francis, and B. Masters. 1992. Mycobacterium haemophilum causing perihilar or cervical lymphadenitis in healthy children. J. Pediatr. 121:202–205. 4. Atlas, R. M., and J. W. Snyder. 2006. Handbook of media for clinical microbiology, 2nd ed., p. 307. CRC Press, Boca Raton, FL. 5. Azov, A. G., J. Koch, and S. J. Hamilton-Dutoit. 2005. Improved diagnosis of mycobacterial infections in formalin-fixed and paraffin-embedded sections with nested polymerase chain reaction. APMIS 113:586–593. 6. Bachmann, S., U. Schnyder, G. E. Pfyffer, R. Lu ¨thy, and R. Weber. 1996. Mycobacterium haemophilum infection in a patient with AIDS. Dtsch. Med. Wochenschr. 121:1189–1192. 7. Reference deleted. 8. Bekou, V., A. Bu ¨chau, M. J. Flaig, T. Ruzicka, and M. Hogardt. 2011. Cutaneous infection by Mycobacterium haemophilum and kansasii in an IgA-deficient man. BMC Dermatol. 11:3. 9. Bernard, E. M., F. F. Edwards, T. E. Kiehn, S. T. Brown, and D. Armstrong. 1993. Activities of antimicrobial agents against clinical isolates of Mycobacterium haemophilum. Antimicrob. Agents Chemother. 37:2323–2326. 10. Besra, G. S., et al. 1991. Structural elucidation and antigenicity of a novel glycolipid antigen from Mycobacterium haemophilum. Biochemistry 30: 7772–7777. 11. Bosma, F., et al. 2004. Mycobacterium reverse hybridization line-probe assay used to diagnose disseminated Mycobacterium haemophilum infection in a child with acute lymphoblastic leukemia. Eur. J. Clin. Microbiol. Infect. Dis. 23:345–347. 12. Branger, B., et al. 1985. Mycobacterium haemophilum and Mycobacterium xenopi associated infection in a renal transplant patient. Clin. Nephrol. 23:46–49. 13. Bruijnesteijn van Coppenraet, E. S., et al. 2004. Real-time PCR assay using fine-needle aspirates and tissue biopsy specimens for rapid diagnosis of mycobacterial lymphadenitis in children. J. Clin. Microbiol. 42:2644–2650. 14. Bruijnesteijn van Coppenraet, L. E. S., E. J. Kuijper, J. A. Lindeboom, J. M. Prins, and E. C. Claas. 2005. Mycobacterium haemophilum and lymphadenitis in children. Emerg. Infect. Dis. 11:62–68. 15. Bruijnesteijn van Coppenraet, L. E. S., V. T. H. B. M. Smit, K. E. Templeton, E. C. J. Claas, and E. J. Kuijper. 2007. Application of real-time PCR to recognize atypical mycobacteria in archival skin biopsies: high prevalence of Mycobacterium haemophilum. Diagn. Mol. Pathol. 16:81–86. 16. Bruijnesteijn van Coppenraet, L. E. S., et al. 2009. Amplified fragment length polymorphism analysis of human clinical isolates of Mycobacterium haemophilum from different continents. Clin. Microbiol. Infect. 15:924–930. 17. Buijtels, P. C., and P. L. Petit. 2005. Comparison of NaOH-N-acetyl cysteine and sulfuric acid decontamination methods for recovery of mycobacteria from clinical specimens. J. Microbiol. Methods 62:83–88. 18. Buijtels, P. C., et al. 2008. Rapid identification of mycobacteria by Raman spectroscopy. J. Clin. Microbiol. 46:961–965. 19. Busam, K. J., T. E. Kiehn, S. P. Salob, and P. L. Myskowski. 1999. Histologic reactions to cutaneous infections by Mycobacterium haemophilum. Am. J. Surg. Pathol. 23:1379–1385. 20. Cameselle, D., et al. 2007. Sporotrichoid cutaneous infection by Mycobacterium haemophilum in an AIDS patient. Actas Dermosifiliogr. 98:188–193. (In Spanish.) 21. Campbell, L. B., M. Maroon, H. Pride, D. C. Adams, and W. B. Tyler. 2006.

CLIN. MICROBIOL. REV. Mycobacterium haemophilum in an immunosuppressed child. Pediatr. Dermatol. 23:481–483. 22. Castro-Silva, A. N., et al. 2011. Cutaneous Mycobacterium haemophilum infection in a kidney transplant recipient after acupuncture treatment. Transpl. Infect. Dis. 13:33–37. 23. Centers for Disease Control. 1991. Mycobacterium haemophilum infections—New York City metropolitan area, 1990–1991. MMWR Morb. Mortal. Wkly. Rep. 40:636–637, 643. 24. Chang, C. T., L. Y. Wang, C. Y. Liao, and S. P. Huang. 2002. Identification of nontuberculous mycobacteria existing in tap water by PCR-restriction fragment length polymorphism. Appl. Environ. Microbiol. 68:3159–3161. 24a.CLSI. 2005. Susceptibility testing of mycobacteria, nocardiae, and other aerobic actinomycetes; approved standard, 2nd ed. M24-A2. CLSI, Wayne, PA. 25. Cohen, Y. H., et al. 2008. Mycobacterium haemophilum and lymphadenitis in immunocompetent children, Israel. Emerg. Infect. Dis. 14:1437–1439. 26. Cooper, D. K. C., et al. 1983. Infectious complications after heart transplantation. Thorax 38:822–828. 27. Covert, T. C., M. R. Rodgers, A. L. Reyes, and G. N. Stelma, Jr. 1999. Occurrence of nontuberculous mycobacteria in environmental samples. Appl. Environ. Microbiol. 65:2492–2496. 28. Da Mata, O., et al. 2008. The diagnosis of two cases of cutaneous ulcer caused by infection with Mycobacterium haemophilum: direct identification in a clinical sample by polymerase chain reaction-restriction endonuclease analysis. Int. J. Dermatol. 47:820–823. 29. Darling, T. N., et al. 1994. Treatment of Mycobacterium haemophilum infection with an antibiotic regimen including clarithromycin. Br. J. Dermatol. 131:376–379. 30. Davis, B. R., J. Brumbach, W. J. Sanders, and E. Wolinsky. 1982. Skin lesions caused by Mycobacterium haemophilum. Ann. Intern. Med. 97:723– 724. 31. Dawson, D. J., Z. M. Blacklock, and D. W. Kane. 1981. Mycobacterium haemophilum causing lymphadenitis in an otherwise healthy child. Med. J. Aust. 2:289–290. 32. Dawson, D. J., and F. F. Jennis. 1980. Mycobacteria with a growth requirement for ferric ammonium citrate, identified as Mycobacterium haemophilum. J. Clin. Microbiol. 11:190–192. 33. Dever, L. L., J. W. Martin, B. Seaworth, and J. H. Jorgensen. 1992. Varied presentations and responses to treatment of infections caused by Mycobacterium haemophilum in patients with AIDS. Clin. Infect. Dis. 14:1195–1200. 34. Devulder, G., M. Pe´rouse de Montclos, and J. P. Flandrois. 2005. A multigene approach to phylogenetic analysis using the genus Mycobacterium as a model. Int. J. Syst. Evol. Microbiol. 55:293–302. 35. Dobos, K. M., F. D. Quinn, D. A. Ashford, C. R. Horsburgh, and C. H. King. 1999. Emergence of a unique group of necrotizing mycobacterial diseases. Emerg. Infect. Dis. 5:367–378. 36. Dore, N., J. P. Collins, and E. Mankiewicz. 1979. A sporotrichoid-like Mycobacterium kansasii infection of the skin treated with minocycline hydrochloride. Br. J. Dermatol. 101:75–79. 37. Elsayed, S., and R. Read. 2006. Mycobacterium haemophilum osteomyelitis: case report and review of the literature. BMC Infect. Dis. 6:70. 38. Endo, T., et al. 2001. Mycobacterium haemophilum infection in a Japanese patient with AIDS. J. Infect. Chemother. 7:186–190. 39. Fairhurst, R. M., et al. 2002. Mycobacterium haemophilum infections in heart transplant recipients: case report and review of the literature. Am. J. Transplant. 2:476–479. 40. Falkinham, J. O., III, C. D. Norton, and M. W. LeChevallier. 2001. Factors influencing numbers of Mycobacterium avium, Mycobacterium intracellulare, and other mycobacteria in drinking water distribution systems. Appl. Environ. Microbiol. 67:1225–1231. 41. Falkinham, J. O., III. 2009. Surrounded by mycobacteria: nontuberculous mycobacteria in the human environment. J. Appl. Microbiol. 107:356–367. 42. Feldman, R. A., and E. Hershfield. 1974. Mycobacterial skin infection by an unidentified species: a report of 29 patients. Ann. Intern. Med. 80:445–452. 43. Fischer, L. J., F. D. Quinn, E. H. White, and C. H. King. 1996. Intracellular growth and cytotoxicity of Mycobacterium haemophilum in a human epithelial cell line (Hec-1-B). Infect. Immun. 64:269–276. 44. Fischer, L. J., F. D. Quinn, L. Kikuta-Oshima, E. M. Ribot, and C. H. King. 1996. Identification of genes specifically expressed by Mycobacterium haemophilum in association with human epithelial cells. Ann. N. Y. Acad. Sci. 797:277–279. 45. Fordham von Reyn, C., et al. 1994. Dual skin testing with Mycobacterium avium sensitin and purified protein derivate: an open study of patients with M. avium complex infection or tuberculosis. Clin. Infect. Dis. 19:15–20. 46. Fordham von Reyn, C., et al. 1998. Dual skin testing with Mycobacterium avium sensitin and purified protein derivate to discriminate pulmonary disease due to M. avium complex from pulmonary disease due to Mycobacterium tuberculosis. J. Infect. Dis. 177:730–736. 47. Friedli, A., J. Krischer, B. Hirschel, J. H. Saurat, and M. Peche`re. 2000. An annular plaque due to Mycobacterium haemophilum infection in a patient with AIDS. J. Am. Acad. Dermatol. 43:913–915. 48. Gardy, J. L., et al. 2011. Whole-genome sequencing and social-network analysis of a tuberculosis outbreak. N. Engl. J. Med. 364:730–739.

VOL. 24, 2011 49. Geisler, W. M., R. D. Harrington, C. K. Wallis, J. P. Harnisch, and W. C. Liles. 2002. Broad spectrum of dermatologic manifestations caused by Mycobacterium haemophilum infection. Arch. Dermatol. 138:229–230. 50. Gey van Pittius, N. C., et al. 2006. Evolution and expansion of the Mycobacterium tuberculosis PE and PPE multigene families and their association with the duplication of the ESAT-6 (esx) gene cluster regions. BMC Evol. Biol. 6:95. 51. Gibson, M. K., S. A. Villano, C. A. Huff, and J. Cofrancesco. 1999. Recrudescence of cutaneous Mycobacterium haemophilum lesions following tetanus immunization. Clin. Infect. Dis. 28:919–920. 52. Giulieri, S., et al. 2011. Outbreak of Mycobacterium haemophilum infections after permanent makeup of the eyebrows. Clin. Infect. Dis. 52:488–491. 53. Goodman, M. D., G. N. Elsasser, and J. Biehle. 2004. Cutaneous ulceration in a patient with HIV. Am. Fam. Physician 70:1537–1538. 54. Gouby, A., B. Branger, R. Oules, and M. Ramuz. 1988. Two cases of Mycobacterium haemophilum infection in a renal-dialysis unit. J. Med. Microbiol. 25:299–300. 55. Griffith, D. E., et al. 2007. An official ATS/IDSA statement: diagnosis, treatment, and prevention of nontuberculous mycobacterial diseases. Am. J. Respir. Crit. Care Med. 175:367–416. 56. Gruschke, A., R. Enzensberger, and V. Brade. 2002. Osteomyelitis of the tibial head caused by Mycobacterium haemophilium in a patient with AIDS. Dtsch. Med. Wochenschr. 127:1947–1950. 57. Gupta, I., et al. 1992. Mycobacterium haemophilum osteomyelitis in an AIDS patient. N. J. Med. 89:201–202. 58. Gutierrez, M. C., et al. 2005. Ancient origin and gene mosaicism of the progenitor of Mycobacterium tuberculosis. PLoS Pathog. 1:e5. 59. Haimi-Cohen, Y., A. Zeharia, M. Mimouni, M. Soukhaman, and J. Amir. 2001. Skin indurations in response to tuberculin testing in patients with nontuberculous mycobacterial lymphadenitis. Clin. Infect. Dis. 33:1786– 1788. 60. Haimi-Cohen, Y., et al. 2009. Chronic cheek lesions: an unusual manifestation of nontuberculous mycobacterial cervicofacial infection. J. Pediatr. 155:746–748. 61. Hanau, L. H., A. Leaf, R. Soeiro, L. M. Weiss, and S. S. Pollack. 1994. Mycobacterium marinum infection in a patient with the acquired immunodeficiency syndrome. Cutis 54:103–105. 62. Harmsen, D., et al. 2003. RIDOM: comprehensive and public sequence database for identification of Mycobacterium species. BMC Infect. Dis. 3:26. 63. Hernandez-Divers, S. J., and D. Shearer. 2002. Pulmonary mycobacteriosis caused by Mycobacterium haemophilum and M. marinum in a royal python. J. Am. Vet. Med. Assoc. 220:1661–1663. 64. Hirsch, R., S. M. Miller, S. Kazi, T. R. Cate, and J. D. Reveille. 1996. Human immunodeficiency virus-associated atypical mycobacterial skeletal infections. Semin. Arthritis Rheum. 25:347–356. 65. Holladay, K. L., and J. K. Carmichael. 1996. Mycobacterium haemophilum cellulitis and osteomyelitis in a man with AIDS. J. Am. Board Fam. Pract. 9:122–124. 66. Holton, J., P. Nye, and R. Miller. 1991. Mycobacterium haemophilum infection in a patient with AIDS. J. Infect. 23:303–306. 67. Hsiao, C. H., Y. T. Lin, C. C. Lai, C. H. Chou, and P. R. Hsueh. 2010. Identification of nontuberculous mycobacterial infection by IS6110 and hsp65 gene analysis on lung tissues. Diagn. Microbiol. Infect. Dis. 68:241– 246. 68. Hussein, Z., O. Landt, B. Wirths, and N. Wellinghausen. 2009. Detection of non-tuberculous mycobacteria in hospital water by culture and molecular methods. Int. J. Med. Microbiol. 299:281–290. 69. Jacob, B., B. M. Debey, and D. Bradway. 2006. Spinal intradural Mycobacterium haemophilum granuloma in an American bison (Bison bison). Vet. Pathol. 43:998–1000. 70. Jang, E. Y., et al. 2007. Case of pyomyositis due to Mycobacterium haemophilum in a renal transplant recipient. J. Clin. Microbiol. 45:3847–3849. 71. Julia ´n, E., et al. 2010. Microscopic cords, a virulence-related characteristic of Mycobacterium tuberculosis, are also present in nonpathogenic mycobacteria. J. Bacteriol. 192:1751–1760. 72. Kamboj, M., et al. 2008. Mycobacterium haemophilum infection after alemtuzumab treatment. Emerg. Infect. Dis. 14:1821–1823. 73. Keller, M., A. Mak, L. Thibert, P. Rene, and M. B. Klein. 2008. Mycobacterium haemophilum epididymal abscess in a renal transplant patient. J. Clin. Microbiol. 46:2459–2460. 74. Kent, M. L., et al. 2004. Mycobacteriosis in zebrafish (Danio rerio) research facilities. Comp. Biochem. Physiol. C Toxicol. Pharmacol. 138:383–390. 75. Kiehn, T. E., and M. White. 1994. Mycobacterium haemophilum: an emerging pathogen. Eur. J. Clin. Microbiol. Infect. Dis. 13:925–931. 76. Kiehn, T. E., et al. 1993. A cluster of four cases of Mycobacterium haemophilum infection. Eur. J. Clin. Microbiol. Infect. Dis. 12:114–118. 77. Kikuchi, K., E. M. Bernard, T. E. Kiehn, D. Armstrong, and L. W. Riley. 1994. Restriction fragment length polymorphism analysis of clinical isolates of Mycobacterium haemophilum. J. Clin. Microbiol. 32:1763–1767. 78. Klein, J. L., T. J. Brown, and G. L. French. 2001. Rifampin resistance in Mycobacterium kansasii is associated with rpoB mutations. Antimicrob. Agents Chemother. 45:3056–3058.

MYCOBACTERIUM HAEMOPHILUM INFECTIONS

715

79. Kristjansson, M., V. M. Bieluch, and P. D. Byeff. 1991. Mycobacterium haemophilum infection in immunocompromised patients: case report and review of the literature. Rev. Infect. Dis. 13:906–910. 80. Kuijper, E. J., F. W. Wit, J. Veenstra, and E. C. Bo ¨ttger. 1997. Recovery of Mycobacterium haemophilum skin infection in an HIV-1-infected patient after the start of antiretroviral triple therapy. Clin. Microbiol. Infect. 3:584– 585. 81. Lederman, C., et al. 1994. Mycobacterium haemophilum cellulites in a heart transplant recipient. J. Am. Acad. Dermatol. 30:804–805. 82. Lee, W. J., et al. 2010. Non-tuberculous mycobacterial infections of the skin: a retrospective study of 29 cases. J. Dermatol. 37:965–972. 83. Lefkowitz, R. A., and R. D. Singson. 1998. Considering Mycobacterium haemophilum in the differential diagnosis for lytic bone lesions in AIDS patients who present with ulcerating skin lesions. Skeletal Radiol. 27:334– 336. 84. Lerner, G. W., A. Safdar, and S. Coppel. 1995. Mycobacterium haemophilum infection in AIDS. Infect. Dis. Clin. Pract. 4:233–236. 85. Lin, J. H., W. Chen, J. Y. Lee, J. J. Yan, and J. J. Huang. 2003. Disseminated cutaneous Mycobacterium haemophilum infection with severe hypercalcaemia in a failed renal transplant recipient. Br. J. Dermatol. 149:200– 202. 86. Lindeboom, J. A. 2011. Conservative wait-and-see therapy versus antibiotic treatment for nontuberculous mycobacterial cervicofacial lymphadenitis in children. Clin. Infect. Dis. 52:180–184. 87. Lindeboom, J. A., et al. 2009. Esthetic outcome of surgical excision versus antibiotic therapy for nontuberculous mycobacterial cervicofacial lymphadenitis in children. Pediatr. Infect. Dis. J. 28:1028–1030. 88. Lindeboom, J. A., E. J. Kuijper, E. S. Bruijnesteijn van Coppenraet, R. Lindeboom, and J. M. Prins. 2007. Surgical excision versus antibiotic treatment for nontuberculous mycobacterial cervicofacial lymphadenitis in children: a multicenter, randomized, controlled trial. Clin. Infect. Dis. 44:1057– 1064. 89. Lindeboom, J. A., C. F. Kuijper, and M. van Furth. 2007. Inguinal lymphadenitis caused by Mycobacterium haemophilum in an immunocompetent child. Pediatr. Infect. Dis. J. 26:84–86. 90. Lindeboom, J. A., J. M. Prins, E. S. Bruijnesteijn van Coppenraet, R. Lindeboom, and E. J. Kuijper. 2005. Cervicofacial lymphadenitis in children caused by Mycobacterium haemophilum. Clin. Infect. Dis. 41:1569– 1575. 91. Lindeboom, J. A., E. J. Kuijper, J. M. Prins, L. E. S. Bruijnesteijn van Coppenraet, and R. Lindeboom. 2006. Tuberculin skin testing is useful in the screening for nontuberculous mycobacterial cervicofacial lymphadenitis in children. Clin. Infect. Dis. 43:1547–1551. 92. Lindeboom, J. A., E. J. Kuijper, E. S. Bruijnesteijn van Coppenraet, and J. M. Prins. 2006. First case of an oculofacial lesion due to Mycobacterium haemophilum infection in an immunocompetent child. Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod. 101:774–776. 93. Lomvardias, S., and G. E. Madge. 1972. Chaetoconidium and atypical acid fast bacilli in skin ulcers. Arch. Dermatol. 106:875–876. 94. Lott, J. P., V. P. Werth, and C. L. Kovarik. 2008. Cutaneous Mycobacterium haemophilum infection in iatrogenically immunocompromised patients without transplantation. J. Am. Acad. Dermatol. 59:139–142. 95. Lotz, A., et al. 2010. Rapid identification of mycobacterial whole cells in solid and liquid culture media by matrix-assisted laser desorption ionization–time of flight mass spectrometry. J. Clin. Microbiol. 48:4481–4486. 96. Males, B. M., T. E. West, and W. R. Bartholomew. 1987. Mycobacterium haemophilum infection in a patient with acquired immune deficiency syndrome. J. Clin. Microbiol. 25:186–190. 97. Malouf, M. A., and A. R. Glanville. 1999. The spectrum of mycobacterial infection after lung transplantation. Am. J. Respir. Crit. Care Med. 160: 1611–1616. 98. Martinelli, C., et al. 2004. First case of Mycobacterium haemophilum infection in an AIDS patient in Italy. J. Eur. Acad. Dermatol. Venereol. 18:83–85. 99. McBride, M. E., et al. 1991. Diagnostic and therapeutic considerations for cutaneous Mycobacterium haemophilum infections. Arch. Dermatol. 127: 276–277. 100. McGovern, J., B. C. Bix, and G. W. Webster. 1994. Mycobacterium haemophilum skin disease successfully treated with excision. J. Am. Acad. Dermatol. 30:269–270. 101. Mezo, A., et al. 1979. Unusual mycobacteria in 5 cases of opportunistic infections. Pathology 11:377–384. 102. Millar, M. J., C. Bulliard, C. Balachandran, and A. J. Maloof. 2007. Mycobacterium haemophilum infection presenting as filamentary keratopathy in an immunocompromised adult. Cornea 26:764–766. 103. Minani, T. J., M. A. Saubolle, E. Yu, and Z. Sussland. 2010. Mycobacterium haemophilum as a novel etiology of cervical lymphadenitis in an otherwise healthy adult patient. J. Clin. Microbiol. 48:2636–2639. 104. Modi, D., et al. 2007. Mycobacterium haemophilum: a rare cause of endophthalmitis. Retina 27:1148–1151. 105. Moulsdale, M. T., J. M. Harper, G. N. Thatcher, and B. L. Dunn. 1983.

716

106. 107.

108.

109.

110.

111.

112.

113.

114.

115.

116. 117.

118.

119.

120.

121. 122.

123.

124.

125.

126.

127.

128.

129.

130.

131.

LINDEBOOM ET AL. Infection by Mycobacterium haemophilum, a metabolically fastidious acidfast bacillus. Tubercle 64:29–36. Murdoch, M. E., and I. M. Leigh. 1989. Sporotrichoid spread of cutaneous Mycobacterium chelonei infection. Clin. Exp. Dermatol. 14:309–312. Olsen, R. J., P. L. Cernoch, and G. A. Land. 2006. Mycobacterial synovitis caused by slow-growing nonchromogenic species: eighteen cases and a review of the literature. Arch. Pathol. Lab. Med. 130:783–791. Paech, V., et al. 2002. Remission of cutaneous Mycobacterium haemophilum infection as a result of antiretroviral therapy in a human immunodeficiency virus-infected patient. Clin. Infect. Dis. 34:1017–1019. Pai, H. H., W. C. Chen, and C. F. Peng. 2003. Isolation of non-tuberculous mycobacteria from hospital cockroaches (Periplaneta americana). J. Hosp. Infect. 53:224–228. Pfister, P., et al. 2004. The structural basis of macrolide-ribosome binding assessed using mutagenesis of 23S rRNA positions 2058 and 2059. J. Mol. Biol. 342:1569–1581. Phowthongkum, P., A. Puengchitprapai, N. Udomsantisook, S. Tumwasorn, and C. Suankratay. 2008. Spindle cell pseudotumor of the brain associated with Mycobacterium haemophilum and Mycobacterium simiae mixed infection in a patient with AIDS: the first case report. Int. J. Infect. Dis. 12:421–424. Plemmons, R. M., C. K. McAllister, M. C. Garces, and R. L. Ward. 1997. Osteomyelitis due to Mycobacterium haemophilum in a cardiac transplant patient: case report and analysis of interactions among clarithromycin, rifampin, and cyclosporine. Clin. Infect. Dis. 24:995–997. Pryor, M., et al. 2004. Investigation of opportunistic pathogens in municipal drinking water under different supply and treatment regimes. Water. Sci. Technol. 50(1):83–90. Rajpara, A., J. Sri, and M. Driscoll. 2010. Mycobacterium haemophilum: cutaneous nodules in a renal transplant patient. Dermatol. Online J. 16(7):3. Richter, E., S. Ru ¨sch-Gerdes, and D. Hillemann. 2006. Evaluation of the GenoType Mycobacterium Assay for identification of mycobacterial species from cultures. J. Clin. Microbiol. 44:1769–1775. Ringuet, H., et al. 1999. hsp65 sequencing for identification of rapidly growing mycobacteria. J. Clin. Microbiol. 37:852–857. Rogers, P. L., et al. 1988. Disseminated Mycobacterium haemophilum infection in two patients with the acquired immunodeficiency syndrome. Am. J. Med. 84:640–642. Roth, A., et al. 2000. Novel diagnostic algorithm for identification of mycobacteria using genus-specific amplification of the 16S-23S rRNA gene spacer and restriction endonucleases. J. Clin. Microbiol. 38:1094–1104. Sagi, L., et al. 2010. Mycobacterium haemophilum infection presenting as bilateral cellulitis and annular lesion in a heart transplant recipient. Isr. Med. Assoc. J. 12:57–58. Saleeb, P. G., S. K. Drake, P. R. Murray, and A. M. Zelazny. 2011. Identification of mycobacteria in solid-culture media by matrix-assisted laser desorption ionization–time of flight mass spectrometry. J. Clin. Microbiol. 49:1790–1794. Sampaio, J. L., et al. 2002. Mycobacterium haemophilum: emerging or underdiagnosed in Brazil? Emerg. Infect. Dis. 8:1359–1360. Samra, Z., et al. 1999. Optimal detection and identification of Mycobacterium haemophilum in specimens from pediatric patients with cervical lymphadenopathy. J. Clin. Microbiol. 37:832–834. Saubolle, M. A., T. E. Kiehn, M. H. White, M. F. Rudinsky, and D. Armstrong. 1996. Mycobacterium haemophilum: microbiology and expanding clinical and geographic spectra of disease in humans. Clin. Microbiol. Rev. 9:435–447. Schumacher, O., J. Dabernig, I. Nenadic, G. Ingianni, and C. Cedidi. 2008. Severe course of a rare non-tuberculous mycobacteriosis (M. haemophilum) of the hand—case report and strategic comments. Handchir. Mikrochir. Plast. Chir. 40:342–347. Seitz, C. S., A. Trautemann, E. B. Bro ¨cker, M. Abele-Horn, and M. Goebeler. 2008. Skin nodules in rheumatoid arthritis due to infection with Mycobacterium haemophilum. Acta Derm. Venereol. 88:428–429. Shah, M. K., A. Sebti, T. E. Kiehn, S. A. Massarella, and K. A. Sepkowitz. 2001. Mycobacterium haemophilum in immunocompromised patients. Clin. Infect. Dis. 33:330–337. Shih, J. Y., et al. 1998. Pyomyositis due to Mycobacterium haemophilum in a patient with polymyositis and long-term steroid use. Clin. Infect. Dis. 26:505–507. Smith, S., G. D. Taylor, and E. A. Fanning. 2003. Chronic cutaneous Mycobacterium haemophilum infection acquired from coral injury. Clin. Infect. Dis. 37:100–101. Somosko ¨vi, A., and P. Magyar. 1999. Comparison of the Mycobacteria Growth Indicator Tube with MB Redox, Lo ¨wenstein-Jensen, and Middlebrook 7H11 media for recovery of mycobacteria in clinical specimens. J. Clin. Microbiol. 37:1366–1369. Sompolinsky, D., A. Lagziel, D. Naveh, and T. Yankilevitz. 1978. Mycobacterium haemophilum sp. nov., a new pathogen of humans. Int. J. Syst. Bacteriol. 28:67–75. Sompolinsky, D., A. Lagziel, and I. Rosenberg. 1979. Further studies of a

CLIN. MICROBIOL. REV.

132.

133. 134.

135.

136. 137. 138.

139.

140.

141. 142.

143.

144.

145.

146.

147.

148.

149. 150. 151.

152.

153.

154.

155. 156.

157.

158.

159.

new pathogenic mycobacterium (M. haemophilum sp. nov.). Can. J. Microbiol. 25:217–226. Soubani, A. O., A. Mohammed, and S. Forlenza. 1994. Successful treatment of disseminated Mycobacterium haemophilum infection in a patient with AIDS. Clin. Infect. Dis. 18:475. Sowden, D., R. Kemp, and D. Dawson. 1993. Osteomyelitis due to Mycobacterium haemophilum in a patient with AIDS. Pathology 25:308–309. Stauffer, F., et al. 1998. Genus level identification of mycobacteria from clinical specimens by using an easy-to-handle Mycobacterium-specific PCR assay. J. Clin. Microbiol. 36:614–617. Straus, W. L., et al. 1994. Clinical and epidemiologic characteristics of Mycobacterium haemophilum, an emerging pathogen in immunocompromised patients. Ann. Intern. Med. 120:118–125. Stu ¨renburg, E. E., et al. 2004. Disseminated Mycobacterium haemophilum infection as initial manifestation of AIDS. Tuberculosis 84:341–345. Swart, R. M., et al. 2009. Nontuberculous mycobacteria infection and tumor necrosis factor-alpha antagonists. Emerg. Infect. Dis. 15:1700–1701. Tan, H. H., A. Tan, C. Theng, and S. K. Ng. 2004. Cutaneous Mycobacterium haemophilum infections in immunocompromised patients in a dermatology clinic in Singapore. Ann. Acad. Med. Singapore 33:532–536. Teh, C. L., K. O. Kong, A. P. Chong, and H. Badsha. 2002. Mycobacterium haemophilum infection in an SLE patient on mycophenolate mofetil. Lupus 11:249–252. Thibert, L., and S. Lapierre. 1993. Routine application of high-performance liquid chromatography for identification of mycobacteria. J. Clin. Microbiol. 31:1759–1763. Thibert, L., F. Lebel, and B. Martineau. 1990. Two cases of Mycobacterium haemophilum infection in Canada. J. Clin. Microbiol. 28:621–623. Tobler, N. E., M. Pfunder, K. Herzog, J. E. Frey, and M. Altwegg. 2006. Rapid detection and species identification of Mycobacterium spp. using real-time PCR and DNA-microarray. J. Microbiol. Methods 66:116–124. Tortoli, E., et al. 2001. Performance assessment of new multiplex probe assay for identification of mycobacteria. J. Clin. Microbiol. 39:1079– 1084. Tortoli, E., A. Mariottini, and G. Mazzarelli. 2003. Evaluation of INNOLiPA MYCOBACTERIA v2: improved reverse hybridization multiple DNA probe assay for mycobacterial identification. J. Clin. Microbiol. 41: 4418–4420. Tuohy, M. J., G. S. Hall, M. Sholtis, and G. W. Procop. 2005. Pyrosequencing as a tool for the identification of common isolates of Mycobacterium sp. Diagn. Microbiol. Infect. Dis. 51:245–250. Vaerewijck, M. J., G. Huys, J. C. Palomino, J. Swings, and F. Portaels. 2005. Mycobacteria in drinking water distribution systems: ecology and significance for human health. FEMS Microbiol. Rev. 29:911–934. Van de Griendt, E. J., P. J. Rietra, and R. N. van Andel. 2003. Mycobacterium haemophilum as the cause of lymphadenitis in the neck in an otherwise healthy boy. Ned. Tijdschr. Geneeskd. 147:1367–1369. Van Ingen, J., M. J. Boeree, P. N. Dekhuijzen, and D. van Soolingen. 2008. Mycobacterial disease in patients with rheumatic disease. Nat. Clin. Pract. Rheumatol. 4:649–656. Von Stebut, E., K. Wiest, and W. Braeuninger. 2005. Chronic infiltrates and persisting ulcerations on the arms and legs. Arch. Dermatol. 141:897–902. Walder, B. K., et al. 1976. The skin and immunosuppression. Aust. J. Dermatol. 17:94–97. Wang, S. X., L. H. Sng, H. N. Leong, and B. H. Tan. 2004. Direct identification of Mycobacterium haemophilum in skin lesions of immunocompromised patients by PCR-restriction endonuclease analysis. J. Clin. Microbiol. 42:3336–3338. Ward, M. S., K. V. Lam, P. K. Cannell, and R. P. Herrmann. 1999. Mycobacterial central venous catheter tunnel infection: a difficult problem. Bone Marrow Transplant. 24:325–329. Weinstock, D. M., M. B. Feinstein, K. A. Sepkowitz, and A. Jakubowski. 2003. High rates of infection and colonization by nontuberculous mycobacteria after allogeneic hematopoietic stem cell transplantation. Bone Marrow Transplant. 31:1015–1021. Whipps, C. M., S. T. Dougan, and M. L. Kent. 2007. Mycobacterium haemophilum infections of zebrafish (Danio rerio) in research facilities. FEMS Microbiol. Lett. 270:21–26. White, M. P., H. Bangash, K. M. Goel, and P. A. Jenkins. 1986. Nontuberculous mycobacterial lymphadenitis. Arch. Dis. Child. 61:368–371. White, M. H., E. Papadopoulos, T. N. Small, T. E. Kiehn, and D. Armstrong. 1995. Mycobacterium haemophilum infections in bone marrow transplant recipients. Transplantation 60:957–960. White, D. A., T. E. Kiehn, A. Y. Bondoc, and S. A. Massarella. 1999. Pulmonary nodule due to Mycobacterium haemophilum in an immunocompetent host. Am. J. Respir. Crit. Care Med. 160:1366–1368. Whittier, S., R. L. Hopfer, M. R. Knowles, and P. H. Gilligan. 1993. Improved recovery of mycobacteria from respiratory secretions of patients with cystic fibrosis. J. Clin. Microbiol. 31:861–864. Winter, N., et al. 1995. Characterization of the gene encoding the immunodominant 35 kDa protein of Mycobacterium leprae. Mol. Microbiol. 16: 865–876.

VOL. 24, 2011

MYCOBACTERIUM HAEMOPHILUM INFECTIONS

717

160. Woods, G. L., N. G. Warren, and C. B. Inderlied. 2007. Susceptibility test methods: mycobacteria, nocardia, and other actinomycetes, p. 1223–1248. In P. R. Murray, E. J. Baron, J. H. Jorgensen, M. L. Landry, and M. A. Pfaller (ed.), Manual of clinical microbiology, 9th ed. ASM Press, Washington, DC. 161. Xiong, L., F. Kong, Y. Yang, J. Cheng, and G. L. Gilbert. 2006. Use of PCR and reverse line blot hybridization macroarray based on 16S-23S rRNA gene internal transcribed spacer sequences for rapid identification of 34 Mycobacterium species. J. Clin. Microbiol. 44:3544–3550.

162. Yakrus, M. A., and W. L. Straus. 1994. DNA polymorphisms detected in Mycobacterium haemophilum by pulsed-field gel electrophoresis. J. Clin. Microbiol. 32:1083–1084. 163. Yarrish, R. L., et al. 1992. Osteomyelitis caused by Mycobacterium haemophilum: successful therapy in two patients with AIDS. AIDS 6:557– 561. 164. Zeharia, A., et al. 2008. Management of nontuberculous mycobacteriainduced cervical lymphadenitis with observation alone. Pediatr. Infect. Dis. J. 27:920–922.

Jerome A. Lindeboom, M.D., D.D.S., Ph.D., is an Oral and Maxillofacial surgeon at the Department of Oral and Maxillofacial Surgery at the Academic Medical Center (AMC) in Amsterdam and the Academic Centre for Dentistry Amsterdam (ACTA), The Netherlands. He also works as an Oral and Maxillofacial surgeon at the Department of Oral and Maxillofacial Surgery at the Amstelland Hospital, Amstelveen, The Netherlands, and he is editor of the European Journal of Oral Implantology.

Dick van Soolingen, Ph.D., is the head of the Mycobacteria Reference Laboratory at the National Institute for Public Health and the Environment (RIVM) in Bilthoven, The Netherlands. In addition, he is a professor at the Department of Microbiology and of Pulmonary Diseases, Radboud University Nijmegen Medical Centre/University Lung Centre Dekkerswald, Nijmegen, The Netherlands. He produced multiple articles on the clinical relevance of nontuberculous mycobacteria in recent years.

Lesla E. S. Bruijnesteijn van Coppenraet is a clinical molecular microbiologist at the Department of Medical Microbiology and Infectious Diseases in the Isala Clinics, Zwolle, The Netherlands. She conducted a Ph.D. study with a special interest in Mycobacterium haemophilum at the Leiden University Medical Center, The Netherlands, and wrote her thesis about diagnostics of nontuberculous mycobacteria in 2009.

Eduard J. Kuijper, M.D., Ph.D., medical microbiologist, is the head of the Department of Experimental Microbiology at the Leiden University Medical Center. He introduced molecular biology and mass spectrometry in diagnostics of bacterial diseases and initiated research projects on Mycobacterium haemophilum and Clostridium difficile, with interest in epidemiology and pathogenesis. He is currently also leading a national workgroup on microbiological diagnostics of mycobacterial diseases.

Jan M. Prins is an infectious disease (ID) specialist and Professor of Infectious Diseases at the Department of Infectious Diseases, Tropical Medicine and AIDS, at the Academic Medical Center (AMC) in Amsterdam, The Netherlands. He is head of the Infectious Diseases Fellowship Training Program at the AMC in Amsterdam, and he is chairman of the Dutch Working Party on Antibiotic Policy (SWAB) and chairs the SWAB guideline development committee.

CLINICAL MICROBIOLOGY REVIEWS, Oct. 2011, p. 718–733 0893-8512/11/$12.00 doi:10.1128/CMR.00002-11 Copyright © 2011, American Society for Microbiology. All Rights Reserved.

Vol. 24, No. 4

Food Animals and Antimicrobials: Impacts on Human Health Bonnie M. Marshall1,2 and Stuart B. Levy1,2,3* Alliance for the Prudent Use of Antibiotics, Boston, Massachusetts,1 and Department of Molecular Biology and Microbiology2 and Department of Medicine,3 Tufts University School of Medicine, Boston, Massachusetts INTRODUCTION .......................................................................................................................................................718 ANTIMICROBIAL USE IN ANIMALS: EFFECTS ON ANTIBIOTIC RESISTANCE EMERGENCE.........718 Nontherapeutic Agents and Practices ..................................................................................................................719 Salmonella and the Swann Report ..........................................................................................................................719 Impacts of Nontherapeutic Use ............................................................................................................................719 EFFECTS OF BANNING GROWTH PROMOTANTS IN ANIMAL FEEDS IN EUROPE ............................722 Avoparcin .................................................................................................................................................................722 Virginiamycin and Other Antibiotics ...................................................................................................................723 EVIDENCE FOR ANIMAL-TO-HUMAN SPREAD OF ANTIBIOTIC RESISTANCE ....................................723 Resistance Acquisition through Direct Contact with Animals .........................................................................723 Antibiotic Resistance Transmission through the Food Chain .........................................................................725 Emergence of Resistance in Human Infections ..................................................................................................725 ADDRESSING KNOWLEDGE GAPS: RESERVOIRS OF ANTIBIOTIC RESISTANCE...............................727 CONCLUSIONS .........................................................................................................................................................728 ACKNOWLEDGMENTS ...........................................................................................................................................729 REFERENCES ............................................................................................................................................................729 the environment, options for avoiding that harm should be examined and pursued even if the harm is not yet fully understood or proven” (103). This communication summarizes a large number of studies on the links between antimicrobials used for growth promotion, in particular, as well as other nontherapeutic antimicrobial (NTA) use in animal husbandry and aquaculture, and the emergence of antibiotic-resistant bacteria in humans. The FAAIR Report (Facts about Antibiotics in Animals and the Impact on Resistance) of the Alliance for the Prudent Use of Antibiotics (APUA) cites areas where antibiotic use can be curtailed and proposes several viable recommendations that could be utilized to reduce the burden of resistance genes created by nontherapeutic antibiotic use in animals (22). Lastly, we consider whether knowledge gaps exist that need addressing in order to answer persisting questions that fuel the controversy over NTA use in food animals.

INTRODUCTION For many decades, antibiotic resistance has been recognized as a global health problem. It has now been escalated by major world health organizations to one of the top health challenges facing the 21st century (40, 65). Some of its causes are widely accepted, for example, the overuse and inappropriate use of antibiotics for nonbacterial infections such as colds and other viral infections and inadequate antibiotic stewardship in the clinical arena (109). But the relationship of drug-resistant bacteria in people to antibiotic use in food animals continues to be debated, particularly in the United States (11, 14, 38, 44, 48, 96, 124). Many have delved into this question, producing volumes of direct and indirect evidence linking animal use to antibiotic resistance confronting people. Among these are a number of studies which unequivocally support the concern that use of antibiotics in food animals (particularly nontherapeutic use) impacts the health of people on farms and, more distantly, via the food chain (69, 88, 90, 111). While it was hoped by many that the years of experience following the bans on nontherapeutic use of antimicrobials in Europe would clearly signal an end to this practice, arguments continue, largely along the lines of a cost/benefit ratio and perceived deficits in solid scientific evidence. Action in the United States continues to lag far behind that of the European Union, which has chosen to operate proactively based on the “precautionary principle,” a guiding tenet of public health. This principle states that “when evidence points toward the potential of an activity to cause significant widespread or irreparable harm to public health or

ANTIMICROBIAL USE IN ANIMALS: EFFECTS ON ANTIBIOTIC RESISTANCE EMERGENCE Antimicrobials are delivered to animals for a variety of reasons, including disease treatment, prevention, control, and growth promotion/feed efficiency. Antimicrobial growth promotants (AGPs) were first advocated in the mid-1950s, when it was discovered that small, subtherapeutic quantities of antibiotics such as procaine penicillin and tetracycline (1/10 to 1/100 the amount of a therapeutic dose), delivered to animals in feed, could enhance the feed-to-weight ratio for poultry, swine, and beef cattle (142). For many years, the positive effects of this practice were championed, while the negative consequences went undetected. But microbiologists and infectious disease experts facing antibiotic resistance questioned the possible harm from this use (74, 89, 109, 136). They found that farms using AGPs had more resistant bacteria in the intestinal

* Corresponding author. Mailing address: Tufts University School of Medicine, Department of Molecular Biology and Microbiology, 136 Harrison Ave, M&V 803, Boston, MA 02111. Phone: (617) 636-6764. Fax: (617) 636-0458. E-mail: [email protected]. 718

VOL. 24, 2011

FOOD ANIMALS AND ANTIMICROBIALS: HUMAN HEALTH IMPACTS

floras of the farm workers and farm animals than in those for similar people and animals on farms not using AGPs. A prospective in vivo/in situ study in 1975 was performed to evaluate the effect of introducing low-dose in-feed oxytetracycline as an AGP on the intestinal floras of chickens and farm dwellers (111). The results showed not only colonization of the chickens with tetracycline-resistant and other drug-resistant Escherichia coli strains but also acquisition of resistance in E. coli in the intestinal flora of the farm family. Other studies over the ensuing 3 decades further elucidated the quantitative and qualitative relationships between the practice of in-feed antimicrobials for animals and the mounting problem of hard-to-treat, drug-resistant bacterial infections in humans (83, 162). Nontherapeutic Agents and Practices The chief agricultural NTAs, used extensively in the United States and also used in Europe until the 1970s, include drugs that have likewise been employed widely in human medicine. In the absence of complete, unbiased data, this NTA use in the United States is estimated to be equal to (159) or as much as eight times greater than (67, 117) the quantity administered for therapeutic use. More recently, concerns have arisen over the extensive use of antimicrobials in the burgeoning aquaculture industry, which more than doubled between 1994 and 2004 (36, 84). Eighty to 90 percent of total production occurs in Asia, with 67% occurring in China alone (64). In many parts of the world, fish farming is integrated with sewage or industrial wastewater or with land agriculture, as manure and other agricultural residues are commonly employed in fish feed (123). The overcrowding, unhygienic measures, and other manipulations in this intensive, industrial-scale production act as stressors to the fish and promote an increased use of antibiotic prophylaxis, particularly in the shrimp and carnivorous fish (such as salmon) industries. Moreover, even though the aquaculture use of AGPs in Western Europe and North America has been discontinued, therapeutic treatment of fish generally occurs en masse via inclusion in fish food, which results in exposure of the entire body of water to the antibiotic. The broad application of antibiotics in fish food leads to leaching from unconsumed food and feces into the water and pond sediments, where it not only exerts selective pressures on the sediment and water microflora but also can be washed to more distant sites, exposing wild fish and shellfish to trace antimicrobials (36). In this environment, the role of transduction (infection by bacterial phages) is considered highly important in facilitating lateral gene transfer (71). Sorum suggested that, historically, the transfer and emergence of resistance have occurred faster from aquatic bacteria to humans than from terrestrial animal bacteria to humans (141). In the United States, the total fish industry use of antibiotics was estimated to be 204,000 to 433,000 pounds in the mid1990s (25) (about 2% of the nonmedical use in cattle, swine, and poultry [117]). In much of the world, however, antibiotics are unregulated and used indiscriminately, and use statistics are rarely collected (25, 157). Although the total quantities of antibiotics employed in aquaculture are estimated to be smaller than those used in land animal husbandry, there is much greater use of antibiotic families that are also used in

719

human medicine (Table 1). In Chile, for example, ⬃100 metric tons of quinolones are used annually (10-fold greater than the amount used in human medicine), mostly in aquaculture (35). At least 13 different antimicrobials are reportedly used by farmers along the Thai coast (75). Salmonella and the Swann Report Alarmed by the rise in multidrug-resistant Salmonella in the 1960s, the United Kingdom’s Swann Report of 1969 recognized the possibility that AGPs were contributing largely to the problem of drug-resistant infections (144). It concluded that growth promotion with antibiotics used for human therapy should be banned. The recommendation was implemented first in England and then in other European countries and Canada. The practice continued unchanged, however, in the United States and ultimately also continued in Europe, but with agents that were not used therapeutically in humans. Antibiotics such as bacitracin, avoparcin, bambermycins, virginiamycin, and tylosin gained in popularity as narrower-spectrum substitutes that had a smaller impact on the broad range of gut flora. Unforeseen, however, was the structural relationship between some of these agents and agents used clinically in humans (Table 1). This similarity meant that they shared a single bacterial target and that use of one agent could produce cross-resistance to the other. Impacts of Nontherapeutic Use Therapeutics applied properly for the treatment of individual animals tend to control the emergence and propagation of antimicrobial-resistant strains, in large part due to their relatively short-term application and relatively small numbers of animals treated. The resistant strains which may appear are generally diluted out by the return of normal, drug-susceptible commensal competitors (110). In contrast, any extended antibiotic applications, such as the use of AGPs, which are supplied for continuous, low-dose application, select for increasing resistance to the agent. Their use in large numbers of animals, as in concentrated animal feeding operations (CAFOs), augments the “selection density” of the antibiotic, namely, the number (density) of animals producing resistant bacteria. An ecological imbalance results—one that favors emergence and propagation of large numbers of resistance genes (113). The selection is not linked merely to the total amount of antibiotic used in a particular environment but to how many individuals are consuming the drug. Each animal feeding on an antibiotic becomes a “factory” for the production and subsequent dispersion of antibiotic-resistant bacteria. NTA uses are also clearly linked to the propagation of multidrug resistance (MDR), including resistance against drugs that were never used on the farm (10, 52, 59, 60, 92, 107, 111, 132, 141, 153, 154, 164). The chronic use of a single antibiotic selects for resistance to multiple structurally unrelated antibiotics via linkage of genes on plasmids and transposons (111, 143). Studies on the impact of NTA use on resistance in land food animals have focused primarily on three bacterial genera— Enterococcus, Escherichia, and Campylobacter—and, to a lesser extent, on Salmonella and Clostridium. All of the above may be members of the normal gut flora (commensals) of food animals

Control of swine dysentery

Respiratory disease prevention and treatment in poultry Aquaculture (oral/bath/injection)

Carbadox

Carbomycinb

Therapy for swine colibacillosis and dysentery, prevention of early poultry mortality, turkey egg dip AGP for cattle, poultry, sheep, and rabbits; coccidiosis prevention in poultry and sheep AGP for chickens and swine; therapy for swine dysentery, pneumonia, chicken necrotic enteritis, and respiratory disease Poultry coccidiostat

Gentamicinb

Monensin

Maduramycin

Lincomycin

Lasalocid

Flumequin Furazolidone

b

Ionophores

Ionophores

Lincosamides

Ionophores

Aminoglycosides

Fluoroquinolones Nitrofurans

Amphenicols

Macrolides

Cyclopolypeptides Elfamycins Fluoroquinolones

Amphenicols

Macrolides

Quinoxalines

Phosphoglycolipids

Polypeptides

Glycopeptides

Orthosomysins

Aminopenicillins

Glycopeptides

Antimicrobial class

Coccidia, Gram-positive organisms Coccidia, Gram-positive organisms

Gram-positive organisms

Gram-positive organisms

Gram-positive and -negative organisms

Broad Broad

Broad

Gram-positive organisms

Gram-negative organisms Gram-positive organisms Broad

Broad

Gram-positive organisms

Gram-positive and -negative organisms

Gram-positive organisms

Gram-positive organisms

Gram-positive organisms

Gram-positive organisms

Moderate

Gram-positive organisms

Spectrum of activity

No

No

Rare

No

Yes

No Yes

No

Yes

Yes No No

Yes

No

No

No

Yes (zinc bacitracin)

No

No

Yes

No

Use in human medicine

Not demonstrated

Not demonstrated

Erythromycin and other macrolides and lincosamides, clindamycin

Not demonstrated

Other aminoglycosides

Fluoroquinolones and quinolones

All amphenicols

Oleandomycin and other macrolides and lincosamides

All polymyxins Other elfamycins only All quinolones

All amphenicols

Other macrolides

Other quinoxolines

Vancomycin, teicoplanin

Actinomycin, colistin, polymyxin B

Vancomycin, teicoplanin

Everninomycin

All penicillins

Vancomycin, teicoplanin

Structurally related antibiotic(s)/antibiotic(s) with shared cross-resistance

Withdrawn from EU as bovine AGP but authorized as poultry coccidiostat

Approved in EU and U.S.

Approved in U.S.

Approved in EU and U.S.

Not approved in U.S. Banned in U.S. food animals in 2005

Chloramphenicol approved in U.S. for dogs only Used in Japan Not marketed Banned for use in poultry by FDA in 2005; not approved for aquaculture in U.S.

Withdrawn from EU in 2006; available in U.S. Withdrawn due to worker toxicity in EU and Canada; available in U.S. and Mexico

Withdrawn from EU; not licensed in U.S. Withdrawn from EU in 1997; not licensed in U.S. Withdrawn from EU in 1999; available in U.S.; used in Japan

Withdrawn from EU in 1997; not licensed in U.S.

Commentsc

MARSHALL AND LEVY

Bovine AGP; prevention/control of coccidiosis in bovines, poultry, and goats

Aquaculture (oral/bath/injection); AGP for poultry, cattle, and swine; therapy for poultry respiratory disease and bovine mastitis Respiratory disease treatment of cattle and swine Aquaculture (oral) Aquaculture (oral/bath)

Erythromycinb

Florfenicol

Broiler, swine, and cattle feed AGP for swine Therapy for bovine and swine respiratory disease, use in aquaculture (oral/bath)

Colistin Efrotomycin Enrofloxacinb

Chloramphenicol

Bambermycin

AGP for poultry, beef cattle, and swine; control of swine dysentery and bacterial enteritis; control of poultry enteritis AGP for poultry, swine, and cattle

Bacitracin/zinc bacitracin

Avoparcin

b

AGP

Aquaculture, oral treatment of swine colibacillosis, treatment of bovine bacterial enteritis and subclinical mastitis AGP for broilers

Amoxicillin,b ampicillinb

Avilamycin

Bovine AGP

Purpose

Ardacin

Antibiotic

TABLE 1. Antimicrobials used in food animal productiona

720 CLIN. MICROBIOL. REV.

AGP in poultry and swine

AGP for poultry and swine, poultry coccidiostat, treatment of swine dysentery Swine AGP, prevention/control of swine dysentery and porcine intestinal adenomatosis, control of Clostridium perfringens in growers Swine AGP, treatment of bovine mastitis

Oxolinic acidb Pristinamycin

Procaine penicillinb

Roxarsone

c

b

a

Swine AGP, therapeutic treatment of mastitis AGP for broilers

Streptogramins

Macrolides

Pleuromutilins

Tetracylines

Aminoglycosides Sulfonamides

Macrolides

Ionophores

Arsenicals

Beta-lactams

Quinolones Streptogramins

Diaminopyrimidines

Macrolides

Quinoxalines

Aminocoumarins

Streptothricins

Ionophores

Aminoglycosides

Gram-positive organisms

Gram-positive organisms, mycoplasmas, spirochetes Gram-positive organisms

Broad

Broad Broad

Gram-positive organisms

Gram-positive organisms

Coccidia

Gram-positive organisms

Broad Gram-positive organisms

Broad

Gram-positive organisms

Gram-positive and -negative organisms

Gram-positive organisms

Gram-negative organisms

Coccidia, Gram-positive organisms

Gram-positive and -negative organisms

Based on data from references 7, 32, 34, 53, 84, 96, 98, 133, and 162. Highly important in human medicine or belongs to critically important class of human antimicrobials. EU, European Union.

Virginiamycin

Tylosinb

Tiamulin

Tetracyclines (oxy- and chlor-)b

Streptomycinb Sulfonamides

Spiramycinb

Aquaculture (bath) Aquaculture (sulfamerazine 关oral兴 and sulfadimethoxine 关oral兴), swine AGP (sulfamethazine), chicken AGP (sulfadimethoxine) AGP for poultry, swine, and cattle; treatment and control of multiple livestock diseases; aquaculture (oral/ bath/injection); control of fish and lobster disease Swine AGP; treatment of swine enteritis, dysentery, and pneumonia

Poultry AGP, prevention of fowl cholera and other infections Aquaculture (oral) AGP

Ormetoprim

Salinomycin

Poultry and swine AGP

Treatment of staph infections, treatment and control of fowl cholera, treatment of bovine mastitis Swine AGP, control of swine dysentery/enteritis

AGP for swine and poultry; treatment/ control of swine enteritis and pneumonia; control of mortality from E. coli in turkeys, bovines, swine, sheep, and goats; control of respiratory and other poultry diseases; aquaculture (bath) Poultry feed coccidiostat, prevention of necrotic enteritis in chickens, AGP for cattle Swine AGP

Oleandomycinb

Olaquindox

Novobiocin

Nourseothricin

Narasin

Neomycinb

Yes

No

No

Yes

Yes Yes

Yes

No

No

Yes

No Yes

No

Yes

No

Yes

No

No

Yes

Erythromycin and other macrolides and lincosamides Quinupristin/ dalfopristin and other streptogramins

Tylosin, erythromycin, and other macrolides

All tetracyclines

All aminoglycosides All sulfonamides

Erythromycin and other macrolides and lincosamides

Not demonstrated

Other arsenicals

Other quinolones Other streptogramins (virginiamycin, quinupristin/ dalfopristin) Other beta-lactams

Erythromycin and other macrolides Trimethoprim

Other quinoxolines

None

Not demonstrated

Gentamicin and other aminoglycosides

Used for disease prevention and treatment in chickens outside the U.S. AGP use withdrawn from EU; available in U.S. Withdrawn from EU; available in U.S.

Withdrawn from EU; authorized in U.S.

Sulfamerazine authorized for U.S. aquaculture, but not marketed

AGP use withdrawn from EU in 1999; not approved in U.S.

Withdrawn from EU

Withdrawn in EU; available in U.S.

Not approved in U.S.

Withdrawn due to worker toxicity in EU and Canada; available in U.S.

Withdrawn from EU; never used in U.S.

Approved in U.S.

Approved in U.S.

VOL. 24, 2011 FOOD ANIMALS AND ANTIMICROBIALS: HUMAN HEALTH IMPACTS 721

722

MARSHALL AND LEVY

but possess the potential to become serious human pathogens. The prospective farm study by Levy in 1975 (111) and studies of others in the following decades clearly demonstrated the selective nature of low-dose, nontherapeutic AGPs on both the pathogenic and commensal flora of food animals such as poultry, swine, and cattle (8, 16, 18, 90, 98, 146, 149). Likewise, in the past decade, studies have demonstrated the selective nature of mass treatment with antimicrobials in aquaculture (36, 62, 84). In the latter, studies have focused on Aeromonas pathogens of both fish and humans and the subsequent highfrequency transfer of their resistance plasmids to E. coli and Salmonella (36). Aarestrup and Carstensen found that resistance derived from use of one NTA (tylosin) was not confined to swine gut bacteria only but could cross species and appear in staphylococci isolated from the skin. While the conversion of gut enterococci to erythromycin (a related human therapeutic) resistance occurred rapidly (within 1 week) the skin-derived resistant organism Staphylococcus hyicus appeared more gradually, escalating to a 5-fold increase over 20 days (5). The finding of bacterial cross-resistance between NTAs used in food animals and human drugs was aptly demonstrated with avoparcin (an AGP) and its close relative vancomycin (an important human therapeutic) when vancomycin-resistant enterococci (VRE) emerged as a serious human pathogen. A connecting link between resistance in animals and humans was revealed when Bates et al. found avoparcin- and vancomycincoresistant enterococci in pigs and small animals from two separate farms. Ribotyping methods showed that some of the patterns from farms and sewage exactly matched those of Enterococcus spp. from the hospital (24). The structures of the two drugs are similar: they are both members of the glycopeptide family (24). Since that time, numerous studies have examined the impacts of newer NTAs on the floras of animals. The use of tylosin and virginiamycin in Norwegian swine and poultry led to high prevalences of resistance to both these agents in Enterococcus faecium (75% to 82% for tylosin and 49 to 70% for virginiamycin) (1). Avilamycin resistance, while significantly associated with avilamycin use, has been observed on both exposed and unexposed farms and was significantly higher in isolates from poultry than in those from swine, despite its use in both these species (4). These findings suggest that other selective agents may be present in the environment or that substances related to avilamycin were not recognized. As described above, not only the drug choice and amount but also the number of animals treated can affect the consequence of its use. Other findings suggest that complex ecologic and genetic factors may play a role in perpetuating resistance (63). Resistance (particularly to tetracycline, erythromycin, and ampicillin) has been found inherently in some antibiotic-free animals, (10, 45, 93, 130), suggesting that its emergence is related to other factors, such as diet, animal age, specific farm type, cohort variables, and environmental pressures (26). While Alexander et al. found MDR (tetracycline plus ampicillin resistance) in bacteria in control animals, the strains that emerged after AGP use were not related to these (10). In addition, resistance to tetracycline was higher for a grain-based diet than a silage-based one. Costa et al. found non-AGP-related resis-

CLIN. MICROBIOL. REV.

tances in enterococci, most likely derived from previous flocks, i.e., the farm environment and the feed source appeared to be responsible for the emergence of the unrelated resistances (45). Khachatryan et al. found an MDR phenotype (streptomycin, sulfonamide, and tetracycline [SSuT] resistance phenotype) propagated by oxytetracycline in a feed supplement, but upon removal of the drug, the phenotype appeared to be maintained by some unknown component of the unmedicated feed supplement, possibly one that selects for another gene that is linked to a plasmid bearing the SSuT resistance phenotype (100). The persistence may also relate to the stability of the plasmid in its host and the fact that expression of tetracycline resistance is normally silent until it is induced by tetracycline. Thus, the energy demands exerted on the host by tetracycline resistance are lower. One can conclude that removal of the antibiotic may not lead to rapid loss of the resistant strain or plasmid.

EFFECTS OF BANNING GROWTH PROMOTANTS IN ANIMAL FEEDS IN EUROPE One of the first bans on AGP use was that imposed on tetracycline by the European Common Market in the mid1970s (39). Prior to institution of the ban in the Netherlands (1961 to 1974), Van Leeuwen et al. had tracked a rise in tetracycline-resistant Salmonella spp. Following the ban, however, they observed a decline in tetracycline resistance in both swine and humans (150). More than 10 years have passed since the final 1999 European Union ban, during which a plethora of studies from multiple European countries, Canada, and Taiwan have examined antibiotic use and resistance trends subsequent to the removal of key AGP drugs, especially avoparcin, and the consequences on vancomycin resistance in Enterococcus (7, 15, 17, 21, 29, 30, 76, 85, 97, 102, 107, 121, 148, 150, 156a). Its structural relationship to and cross-resistance with avoparcin render vancomycin a drug of prime interest for determining the impact of avoparcin in triggering and promoting resistance in human infection.

Avoparcin In many European countries, the use of avoparcin as a feed additive led to frequent isolation of VRE from farm animals and healthy ambulatory people (3, 18, 102). Since the emergence of the enterococcus as a major MDR pathogen, vancomycin has evolved as a key therapy, often as the drug of last resort. Following the 1995 ban on avoparcin, several investigators reported a decline in animal VRE. In Denmark, frequencies peaked at 73 to 80% and fell to 5 to 6% (7, 18) in poultry. In Italy, VRE prevalence in poultry carcasses and cuts decreased from 14.6% to 8% within 18 months of the 1997 ban (121), and in Hungary, a 4-year study showed not only a decline in prevalence of VRE among slaughtered cattle, swine, and poultry after removal of avoparcin but also a decrease in vancomycin MICs (97). In surveillance studies both before and after the German ban in 1996, Klare et al. showed a high frequency of VRE in 1994, followed by a very low frequency of just 25% of poultry food products in 1999 (102). Similar de-

VOL. 24, 2011

FOOD ANIMALS AND ANTIMICROBIALS: HUMAN HEALTH IMPACTS

clines were reported in broiler farms following a ban on avoparcin in Taiwan in 2000 (107). A dramatic reduction in human carriage of VRE also followed the ban on avoparcin. Parallel surveillances of the gut floras of healthy ambulatory people showed that VRE colonization in Germany declined from 13% in 1994 to 4% in 1998 (102), and in Belgium, it declined from 5.7% in 1996 to ⬃0.7% in 2001 (68). Virginiamycin and Other Antibiotics Increased virginiamycin use in Danish broilers during the mid-1990s correlated with a rise in resistant E. faecium prevalence, from 27% to ⬃70% (7). Following the ban, resistance declined to 34% in 2000. Likewise, in Denmark, the 1998 ban on the use of tylosin in swine resulted in a decline in erythromycin (a structurally related macrolide) resistance, from 66% to 30% (49). Avilamycin use in 1995 and 1996 increased resistance in broiler E. faecium strains, from 64% to 77%, while declining applications after 1996 lowered the prevalence to 5% in 2000 (7). Some of these studies revealed a genetic linkage between bacterial macrolide and glycopeptide resistances in swine, such that neither resistance declined in prevalence until both avoparcin (a glycopeptide) and tylosin (a macrolide) use was limited. With a reduction in tylosin use, the prevalence of glycopeptide-resistant enterococci fell to 6% and macrolide resistance fell from nearly 90% to 47% in E. faecium and to 28% in Enterococcus faecalis (7). Notably, the first report of transfer of vancomycin resistance from Enterococcus to Staphylococcus aureus was demonstrated in laboratory mice because of its linkage to macrolide resistance on the same plasmid (119). One concern voiced following the banning of NTAs was that the incidence of disease in animals would rise and result in a parallel increase in therapeutic use. This has become the subject of some debate. Some countries encountered rises in necrotic enteritis in chickens and colitis in swine soon after the institution of AGP bans (33, 159). In Norway, an abrupt increase in necrotizing enteritis (NE) in poultry broilers was reported following the removal of avoparcin, with a coincident rise in antibiotic therapy. When the ionophore feed additive narasin was approved, NE declined once again (77). It was concluded that the ban on avoparcin consumption produced a negligible effect on the need for antibiotic therapy (76). Likewise, in Switzerland, Arnold et al. reported a postban increase in overall antibiotic quantities used in swine husbandry but observed a stable therapy intensity (prescribed daily dose) (15). By 2003, total animal use of antibiotics in Denmark, Norway, and Sweden had declined by 36%, 45%, and 69%, respectively (76). The most thorough postban analysis of this phenomenon comes from Denmark. In a careful review of swine disease emergence, animal production, and antibiotic use patterns over the years 1992 to 2008, Aarestrup et al. reported no overall deleterious effects from the ban on finishers and weaners in the years 1998 and 2000, respectively. Despite an increase in total therapeutic antibiotic consumption immediately following the ban, no lasting negative effects were detected on mortality rate, average daily weight gain, or animal production (6). Moreover, even if therapeutic use increased,

723

the numbers of animals treated would be reduced compared to those with growth promotion use, so selection density would be decreased (113). In summary, the in-depth, retrospective analyses in Denmark shed a different perspective on postban concerns over increased therapeutic use. Over time, it appears that the negative after-effects of the ban have waned. As farmers modified their animal husbandry practices to accommodate the loss of banned NTAs, these disease outbreaks became less prominent. Improved immunity and reduced infection rates led to fewer demands for therapeutic antibiotics. Interestingly, recent studies have shown that the original beneficial aspects observed with AGP use (i.e., weight gain and feed efficiency) appear to have diminished, although the results are mixed and depend upon the kind of animals and type of antibiotic involved. Diarra et al. found no effect on body weight or feed intake in poultry from five different AGPs, and feed efficiency was improved with penicillin only (52). In contrast, Dumonceaux et al. reported a significantly increased body weight (10%) and a 7% increase in feed efficiency with the AGP virginiamycin, but only for the first 15 days (55a). In short, improved farming practices and breeding programs, which may include reduced animal density, better hygiene, targeted therapy, and the use of enzymes, prebiotics, probiotics, and vaccines, appear to have at least partially replaced the beneficial aspects of antibiotic growth promoters (27, 158, 160). EVIDENCE FOR ANIMAL-TO-HUMAN SPREAD OF ANTIBIOTIC RESISTANCE Any use of antibiotics will select for drug-resistant bacteria. Among the various uses for antibiotics, low-dose, prolonged courses of antibiotics among food animals create ideal selective pressures for the propagation of resistant strains. Spread of resistance may occur by direct contact or indirectly, through food, water, and animal waste application to farm fields. It can be augmented greatly by the horizontal transfer of genetic elements such as plasmids via bacterial mating (conjugation). We summarize here the evidence for animal-to-human transfer of resistant bacteria on farms using antibiotics for treatment and/or nontherapeutic use. Resistance Acquisition through Direct Contact with Animals Farm and slaughterhouse workers, veterinarians, and those in close contact with farm workers are directly at risk of being colonized or infected with resistant bacteria through close contact with colonized or infected animals (Table 2). Although this limited transmission does not initially appear to pose a population-level health threat, occupational workers and their families provide a conduit for the entry of resistance genes into the community and hospital environments, where further spread into pathogens is possible (118, 155). The majority of studies examining the transmission of antibiotic-resistant bacteria from animals to farm workers document the prevalence of resistance among farmers and their contacts or among farmers before and after the introduction of antibiotics at their workplace. Direct spread of bacteria from animals to people was first reported by Levy et al., who found

Transfer type

Human infection via direct or indirect animal contact

Human colonization via direct or indirect animal contact

Danish swine and chickens

Spanish chickens (slaughtered)

Enterococcus faecium

E. coli

E. coli

Belgian cattle (ill)

Dutch veal calves

MRSA ST398

E. coli, Salmonella enterica (serovar Typhimurium)

Chinese swine and chickens

E. coli

Beef cattle (ground beef) receiving chlortetracycline AGP German swine (ill)

U.S. chickens

E. coli

Salmonella Newport

French swine

S. aureus, Streptococcus spp., E. coli and other enterobacteria

Animal host(s) U.S. chickens

E. coli

Species tracked

Resistance transferred

Ciprofloxacin

Vancomycin

Apramycin, gentamicin

Streptothricin

Ampicillin, carbenicillin, tetracycline

MDR

Apramycin (not used in human medicine)

Gentamicin

Erythromycin, penicillins, nalidixic acid, chloramphenicol, tetracycline, streptomycin, cotrimoxazole

Tetracycline

Evidence

Direct genetic tracking of resistance plasmid from hamburger meat to infected patients Identification of transferable resistance plasmids found only in human gut and UTI bacteria when nourseothricin was used as swine AGP Plasmid-based transfer of aac(3)-IV gene bearing resistance to a drug used only in animals (apramycin) Clonal spread of E. faecium and horizontal transmission of the vanA gene cluster (Tn1546) found between animals and humans Multiple molecular and epidemiological typing modalities demonstrated avian source of resistant E. coli

Following introduction of tetracycline on a farm, resistant E. coli strains with transferable plasmids were found in caretakers’ gut floras, with subsequent spread to the farm family Phenotypic antibiotic resistance was significantly higher in the commensal floras (nasal, pharyngeal, and fecal) of swine farmers than in those of nonfarmers Increase in phenotypic gentamicin resistance in workers through direct contact with chickens receiving gentamicin prophylactically Detection of aac(3)-IV apramycin resistance gene in humans, with 99.3% homology to that in animal strains Human nasal carriage of the mecA gene was strongly associated with (i) greater intensity of animal contact and (ii) the number of MRSA-positive animals; animal carriage was related to animal antibiotic treatment

95

80

42

90

87

78

164

126

16

111

Reference

MARSHALL AND LEVY

Bacteremic hospital patients

Hospital patients with diarrhea

Hospital inpatients

Swine farmers, family members, community members, UTI patients

Salmonella-infected patients with diarrhea

Veal farmers

Farm workers

Poultry workers

Swine farmers

Animal caretakers, farm family

Recipient host(s)

TABLE 2. Key evidence for transfer of antibiotic resistance from animals to humans

724 CLIN. MICROBIOL. REV.

VOL. 24, 2011

FOOD ANIMALS AND ANTIMICROBIALS: HUMAN HEALTH IMPACTS

the same tetracycline-resistant E. coli strains in the gut flora of chicken caretakers as in the chickens receiving tetracyclinelaced feed (112). The observation extended to the farm family as well and showed an increased frequency of tetracyclineresistant and multidrug-resistant E. coli after several months of use of AGP-laden feed. Studies such as this (which examined a variety of antibiotic classes and assorted pathogens) have consistently shown a higher prevalence of resistant gut bacteria among farm workers than in the general public or among workers on farms not using antibiotics (16, 90, 98, 149). While gentamicin is not approved for growth promotion in the United States, it remains the most commonly used antibiotic in broiler production, being employed for prevention of early poultry mortality (115). A revelatory 2007 study found that the risk for carrying gentamicin-resistant E. coli was 32 times higher in poultry workers than in other members of the community: half of all poultry workers were colonized with gentamicin-resistant E. coli, while just 3% of nonpoultry workers were colonized. Moreover, the occupationally exposed population was at significantly greater risk for carriage of multidrug-resistant bacteria (126). New gene-based methods of analysis provide even stronger evidence for the animal origin of bacteria that colonize or infect humans. Homologous relationships between bacterial resistance genes in humans and farm animals have been identified most commonly for food-borne pathogens such as Escherichia coli and Salmonella (see below) but have also been recorded for various species of Enterococcus and for methicillin-resistant Staphylococcus aureus (MRSA). Zhang and colleagues found E. coli strains resistant to apramycin (an antibiotic used in agriculture but not in human medicine) in a study of Chinese farm workers. All farms in the study that used apramycin as an AGP had workers that carried apramycin resistance genes. The same resistance gene, aac(3)-IV, was present in each swine, poultry, and human isolate, with some resistance profiles also matching across species (164). A group of French scientists found the same resistance gene [aac(3)-IV] in cow, pig, and human E. coli strains that bore resistance to apramycin and gentamicin (42). In another study, similar resistance patterns and genes were detected in E. faecalis and E. faecium strains from humans, broilers, and swine in Denmark (2). Lee sampled MRSA isolates from cattle, pigs, chickens, and people in Korea and found that 6 of the 15 animal isolates containing mecA (the gene responsible for methicillin resistance in S. aureus) were identical to human isolates (108). Antibiotic Resistance Transmission through the Food Chain Consumers may be exposed to resistant bacteria via contact with or consumption of animal products—a far-reaching and more complex route of transmission. There is undeniable evidence that foods from many different animal sources and in all stages of processing contain abundant quantities of resistant bacteria and their resistance genes. The rise of antibiotic-resistant bacteria among farm animals and consumer meat and fish products has been well documented (36, 108, 122, 162). Demonstrating whether such reservoirs of resistance pose a risk to humans has been more challenging as a consequence of the complex transmission routes between farms and consumers and the frequent transfer of resistance genes among host bac-

725

teria. Such correlations are becoming more compelling with the advent of molecular techniques which can demonstrate the same gene (or plasmid) in animal or human strains, even if the isolates are of different species. For example, Alexander et al. showed that drug-resistant Escherichia coli was present on beef carcasses after evisceration and after 24 h in the chiller and in ground beef stored for 1 to 8 days (9). Others isolated ciprofloxacin-resistant Campylobacter spp. from 10% to 14% of consumer chicken products (79, 137). MRSA has been reported to be present in 12% of beef, veal, lamb, mutton, pork, turkey, fowl, and game samples purchased in the consumer market in the Netherlands (50), as well as in cattle dairy products in Italy (120). Likewise, extensive antibiotic resistance has been reported for bacteria, including human pathogens, from farmed fish and market shrimp (56, 84, 140). Some of the antibiotic resistance genes identified in food bacteria have also been identified in humans, providing indirect evidence for transfer by food handling and/or consumption. In 2001, Sorensen et al. confirmed the risk of consuming meat products colonized with resistant bacteria, showing that glycopeptide-resistant Enterococcus faecium of animal origin ingested via chicken or pork lasted in human stool for up to 14 days after ingestion (139). Donabedian et al. found overlap in the pulsed-field gel electrophoresis (PFGE) patterns of gentamicin-resistant isolates from humans and pork meat as well as in those of isolates from humans and grocery chicken (55). They identified that when a gene conferring antibiotic resistance was present in food animals, the same gene was present in retail food products from the same species. Most resistant enterococci possessed the same resistance gene, aac(6⬘)-Ieaph(2⬙)-Ia (55). Emergence of Resistance in Human Infections There is likewise powerful evidence that human consumption of food carrying antibiotic-resistant bacteria has resulted, either directly or indirectly, in acquisition of antibiotic-resistant infections (Table 2). In 1985, scientists in Arizona traced an outbreak of multidrug-resistant Salmonella enterica serovar Typhimurium, which included the death of a 72-year-old woman, to consumption of raw milk. Isolates from most patients were identical to the milk isolates, and plasmid analysis showed that all harbored the same resistance plasmid (145). A 1998 S. Typhimurium outbreak in Denmark was caused by strains with nalidixic acid resistance and reduced fluoroquinolone susceptibility. PFGE revealed that a unique resistance pattern was common to Salmonella strains from all patients, two sampled pork isolates, the swine herds of origin, and the slaughterhouse (118). Samples from gentamicin-resistant urinary tract infections (UTIs) and fecal E. coli isolates from humans and food animal sources in China showed that 84.1% of human samples and 75.5% of animal samples contained the aaaC2 gene for gentamicin resistance (86). Johnson et al. used PFGE and random amplified polymorphic DNA (RAPD) profiles of fluoroquinolone-resistant E. coli strains in human blood and fecal samples and in slaughtered chickens to determine that the two were virtually identical to resistant isolates from geographically linked chickens. Drug-susceptible human E. coli strains, how-

726

MARSHALL AND LEVY

ever, were genetically distinct from poultry bacteria, suggesting that the ciprofloxacin-resistant E. coli strains in humans were imported from poultry rather than originating from susceptible human E. coli (94, 95). Other reports demonstrate a broader linkage of resistance genes through the farm-to-fork food chain. A resistance-specifying blaCMY gene was found in all resistant isolates of Salmonella enterica serotype Newport originating from humans, swine, cattle, and poultry. The host plasmid, which conferred resistance to nine or more antimicrobials, was capable of transmission via conjugation to E. coli as well (165). An observed homology between CMY-2 genes in cephalosporin-resistant E. coli and Salmonella suggested that plasmids conferring resistance had moved between the two bacterial species. The authors found higher rates of CMY-2 in strains from animals than in those from humans, supporting an animal origin for the human pathogen (161). A 2000 study found matching PFGE profiles among vancomycin-resistant Enterococcus faecium isolates from hospitalized humans, chickens, and pigs in Denmark. Molecular epidemiology studies have also linked tetracycline resistance genes from Aeromonas pathogens in a hospital effluent to Aeromonas strains from a fish farm (127). These results support the clonal spread of resistant isolates among different populations (80). Chronologic studies of the emergence of resistance across the food chain also strongly imply that reservoirs of resistance among animals may lead to increased resistance in consumers of animal food products. Bertrand et al. chronicled the appearance of the extended-spectrum beta-lactamase (ESBL) gene CTX-M-2 in Salmonella enterica in Belgium. This resistance element was identified first in poultry flocks and then in poultry meat and, finally, human isolates (28). A recent Canadian study also noted a strong correlation between ceftiofur-resistant bacteria (the pathogen Salmonella enterica serovar Heidelberg and the commensal E. coli) from retail chicken and human infections across Canada. The temporary withdrawal of ceftiofur injection from eggs and chicks dramatically reduced resistance in the chicken strains and the human Salmonella isolates, but the trend reversed when the antibiotic use was subsequently resumed (57). In three countries (United States, Spain, and the Netherlands), a close temporal relationship has been documented between the introduction of fluoroquinolone (sarafloxacin and enrofloxacin) therapy in poultry and the emergence of fluoroquinolone-resistant Campylobacter in human infections. An 8to 16-fold increase in resistance frequency was observed—from 0 to 3% prior to introduction to ⬃10% in the United States and the Netherlands and to ⬃50% in Spain—within 1 to 3 years of the licensure (61, 128, 137). In the Netherlands, this frequency closely paralleled an increase in resistant isolates from retail poultry products (61), while the U.S. study used molecular subtyping to demonstrate an association between the clinical human isolates and those from retail chicken products (137). It is now theorized, from molecular and epidemiological tracking, that the resistance determinants found in salmonella outbreaks (strain DT104) in humans and animals in Europe and the United States likely originated in aquaculture farms of the Far East. The transmissible genetic element contains the florfenicol gene (floR) and the tetracycline class G gene, both

CLIN. MICROBIOL. REV.

of which were traced to Vibrio fish pathogens (Vibrio damsel and Vibrio anguillarum, respectively). Both drugs are used extensively in aquaculture (36). In the above examples, the link to nontherapeutic antibiotic use in the farm animals is still circumstantial and largely implied, often because the authors do not report any statistics on farm use of antibiotics. Interpreting these studies is also difficult because of the widespread resistance to some drugs in bacteria of both animals and humans and the ubiquitous nature of resistance genes. Moreover, the same farmer may use antibiotics for both therapeutic and nontherapeutic purposes. The complexities of the modern food chain make it challenging to perform controlled studies that provide unequivocal evidence for a direct link between antibiotic use in animals and the emergence of antibiotic resistance in food-borne bacteria associated with human disease. While this concrete evidence is limited, a small number of studies have been able to link antibiotic-resistant infection in people with bacteria from antibiotic-treated animals. While not necessarily involving NTAs, these studies substantiate the considerable ease with which bacteria in animals move to people. For example, a multidrugresistant Salmonella enterica strain in a 12-year-old Nebraska boy was traced to his father’s calves, which had recently been treated for diarrhea. Isolates from the child and one of the cows were determined to be the same strain of CMY-2-mediated ceftriaxone-resistant S. enterica (69). It is now believed that the 1992 multiresistant Vibrio cholerae epidemic in Latin America was linked to the acquisition of antibiotic-resistant bacteria arising from heavy antibiotic use in the shrimp industry of Ecuador (13, 156). By comparing the plasmid profiles of MDR Salmonella Newport isolates from human and animal sources, Holmberg et al. provided powerful evidence that salmonella infections in 18 persons from 4 Midwestern states were linked directly to the consumption of hamburger meat from cattle fed subtherapeutic chlortetracycline. A plasmid which bore tetracycline and ampicillin resistance genes was present in the organisms causing serious illness in those persons who ate the hamburger meat and who were also consuming penicillin derivatives for other reasons (87). One of the most compelling studies to date is still Hummel’s tracking of the spread of nourseothricin resistance, reported in 1986. In Germany, nourseothricin (a streptogramin antibiotic) was used solely for growth promotion in swine. Resistance to it was rarely found and was never plasmid mediated. Following 2 years of its use as a growth promotant, however, resistance specified by plasmids appeared in E. coli, not only from the treated pigs (33%) but also in manure, river water, food, and the gut floras of farm employees (18%), their family members (17%), and healthy outpatients (16%) and, importantly, in 1% of urinary tract infections (90). Ultimately, the resistance determinant was detected in Salmonella and Shigella strains isolated from human diarrhea cases (146). The movement of antibiotic resistance genes and bacteria from food animals and fish to people—both directly and indirectly—is increasingly reported. While nontherapeutic use of antibiotics is not directly implicated in some of these studies, there is concern that pervasive use of antimicrobials in farming and widespread antimicrobial contamination of the environment in general may be indirectly responsible. For instance,

VOL. 24, 2011

FOOD ANIMALS AND ANTIMICROBIALS: HUMAN HEALTH IMPACTS

within the past 5 years, MRSA and MDR Staphylococcus aureus have been reported in 25 to 50% of swine and veal calves in Europe, Canada, and the United States (51, 78, 101, 114). Graveland et al. noted that this frequency was higher in veal calves fed antibiotics (78). Studies also show that colonization among farmers correlates significantly with MRSA colonization among their livestock (78, 101, 114, 138). In the Netherlands, colonization of swine farmers was found to be more than 760 times greater than that of patients admitted to Dutch hospitals (155). In a study of nasal swabs from veal and veal calf growers, family members, and employees at 102 veal calf farms in the Netherlands, Graveland et al. found that human MRSA sequence type ST398 carriage among the farmers was strongly associated with the degree of animal contact and the frequency of MRSA-colonized animals on the farm. When ⬍20% of calves were carriers, the estimated prevalence in humans was ⬃1%—similar to that in the general public. With ⬎20% carriage in calves, the prevalence in humans was ⬎10% (78). Recently, MRSA ST398 has appeared in the community. A Dutch woman without any known risk factors was admitted to a hospital with endocarditis caused by MRSA ST398, suggesting a community reservoir which passed on to people (58). Voss et al. demonstrated animal-to-human and human-to-human transmission of MRSA between a pig and pig farmer, among the farmer’s family members, and between a nurse and a patient in the hospital. All isolates had identical random amplified polymorphic DNA profiles (155). Examples of similar MRSA strains among animals and people are mounting (82, 108, 147, 151, 152, 163). ADDRESSING KNOWLEDGE GAPS: RESERVOIRS OF ANTIBIOTIC RESISTANCE Historically, considerable attention has been focused on a very small minority of bacterial species that actually cause disease. However, a vast “sea” of seemingly innocuous commensal and environmental bacteria continuously and promiscuously exchange genes, totally unnoticed (116). A staggeringly diverse group of species maintain a large capacity for carrying and mobilizing resistance genes. These bacteria constitute a largely ignored “reservoir” of resistance genes and provide multiple complex pathways by which resistance genes propagated in animals can directly, or more likely indirectly, make their way over time into human pathogens via food, water, and sludge and manure applied as fertilizer. Horizontal (or lateral) gene transfer studies have identified conjugal mating as the most common means of genetic exchange, and there appear to be few barriers that prevent this gene sharing across a multitude of dissimilar genera (104). While colonic bacteria have received much focused study, water environments such as aquaculture, sludge, freshwater, and wastewaters are prime sites for gene exchange but have been examined minimally for their roles as “mixing pots” and transporters of genes from bacteria of antibiotic-fed animals to humans (116). Aside from the already described impacts of NTA use on bacterial resistance, food animal use of NTAs has broad and far-reaching impacts on these environmental bacteria. It is estimated that 75% to 90% of antibiotics used in food animals are excreted, largely unmetabolized, into the environ-

727

ment (43, 105). Antibiotics or resistant bacteria have been detected in farm dust (81), the air currents inside and emanating from swine feeding operations (41, 72, 129), the groundwater associated with feeding operations (31, 37), and the food crops of soils treated with antibiotic-containing manure (54). This leaching into the environment effectively exposes countless environmental organisms to minute quantities of antibiotic— enough to select bacteria with resistance mutations to promote the emergence and transfer of antibiotic resistance genes among diverse bacterial types (104). The potentially huge impact of all these residual antibiotics on the environmental bacteria that are directly or indirectly in contact with humans has scarcely been examined. The multiple pathways and intricacies of gene exchange have so far thwarted attempts to qualitatively or quantitatively track the movement of these genes in vivo, and thus we are left with minimal direct evidence for linking resistance in animals to that in humans. With extensive gene movement between disparate hosts, it is less likely that the same bacterial hosts will be found in animals and humans and more probable that only the resistance genes themselves will be identifiable in the final pathogens that infect humans. Even these may be altered in their journey through multiple intermediate hosts (161) (Fig. 1). Mounting evidence exists in reports of complex gene “cassettes” which accumulate resistance genes and express multidrug resistance (106, 125). A few investigators have undertaken the challenging task of developing mathematical models in order to predict the impacts of NTAs on human disease (12, 19, 20, 46, 91, 99, 134, 135). Models can be very useful in attempting to define the types of diverse data sets that are seen in this field. Some explore the entire “farm-to-fork” transmission process, while others tackle only portions of this extremely complex chain or adopt a novel backwards approach which looks first at human infections and then calculates the fraction that are potentially caused by NTA use in animals. Most models are deliberately simplified and admittedly omit many aspects of transmission and persistence. Moreover, current models are frequently based on multiple assumptions and have been challenged on the basis of certain shortcomings, such as limitation to single pathogens only, the determination of lethality while ignoring morbidity, and dependence on estimates of probabilities (19). Chief among these, however, is the lack of a complete understanding of the contribution made by commensals, which may play an important role in augmenting the link between animals and humans. Some models are driven by findings of dissimilar strains in animals and humans and therefore arrive at very low probabilities for a causal link between the two (47). A finding of dissimilar strains, however, overlooks two possibilities. First, it does not exclude the existence of small subpopulations of homologous strains that have gone undetected within the gut floras of animals. These may have been amplified temporarily by antibiotic selection and transferred their mobile genetic elements in multiple complex pathways. Subsequently, they may have declined to nondetectable levels or merely been outcompeted by other variants. Second, it overlooks dissimilarities that evolve as genes and their hosts migrate in very complex ways through the environment. Figure 1 illustrates the difficulties in tracking a resistance gene, since these genes are frequently captured in bacteria of different species or strains

728

CLIN. MICROBIOL. REV.

MARSHALL AND LEVY

FIG. 1. Several scenarios may present themselves in the genetic transport that occurs as bacteria migrate from animal to human environments. (A) The same host and its indigenous genes in animals are transported unchanged to humans, with a resulting 100% match of the bacterial strain. (B) The genetic structure passes through one or more different hosts, ending in a new host (humans), with a resulting 100% match of DNA. (C) The host and its plasmid-borne genes pass through the environment, picking up gene cassettes en route, with a resulting 100% match for the host only (a) or a low-% match for DNA only (b). In both examples, the plasmid core remains the same.

which no longer resemble the original host. Over time, even the genes themselves may undergo mutations or become entrapped in gene cassettes that alter their genetic landscape. State-of-the-art technology and thoughtful investigation are often necessary to identify and track the actual strains that link animals and humans. These are facets of modeling that have yet to be explored, and obtaining direct evidence for the origins of specific genes can be highly challenging. In general, the weaknesses of present models lie in their simplicity and the lack of crucial knowledge of microbial loads at each stage of the “farm-to-fork” transmission chain. Many of the available studies that examine links between animals and humans suffer from a failure to examine the antibiotic use practices for the farm animals they investigate. More powerful evidence could have accumulated that would aid in modeling efforts if data on the quantities and uses of farm antibiotics had been reported. These oversights are often due to the lack of registries that record and report the utilization of antibiotics on food animal farms. It is widely advocated that surveillance studies of resistance frequencies at all levels of the transmission chain would aid greatly in reducing our knowledge deficits and would help to inform risk management deliberations (23, 34). A number of localized and international surveillance systems exist for the tracking of human pathogens. In the United States, the National Antimicrobial Resistance Monitoring System (NARMS) has become instrumental in the monitoring of resistance trends in pathogens found in food animals, retail meats, and humans (73). However, at the level of commensals, resistance monitoring is still in its infancy. The Reservoirs of Antibiotic Resistance (ROAR) database (www.roarproject .org) is a fledgling endeavor to promote the accumulation of data that specifically focus on commensal and environmental strains as reservoirs of antibiotic resistance genes. With ad-

vances in detection at the genetic level, the potential for tracking the emergence and spread of horizontally transmissible genes is improving rapidly. By capturing geographic, phenotypic, and genotypic data from global isolates from animal, water, plant, and soil sources, the ROAR project documents the abundance, diversity, and distribution of resistance genes and utilizes commensals as “barometers” for the emergence of resistance in human pathogens. CONCLUSIONS Data gaps continue to fuel the debate over the use of NTAs in food animals, particularly regarding the contribution and quantitation of commensal reservoirs of resistance to resistance in human disease. Nonetheless, it has been argued reasonably that such deficits in surveillance or indisputable demonstrations of animal-human linkage should not hinder the implementation of a ban on the use of nontherapeutic antibiotics (23). Food animals produce an immense reservoir of resistance genes that can be regulated effectively and thus help to limit the negative impacts propagated by this one source. In the mathematical model of Smith et al., which specifically evaluates opportunistic infections by members of the commensal flora, such as enterococci, it was concluded that restricting antibiotic use in animals is most effective when antibioticresistant bacteria remain rare. They suggest that the timing of regulation is critical and that the optimum time for regulating animal antibiotic use is before the resistance problem arises in human medicine (134). A ban on nontherapeutic antibiotic use not only would help to limit additional damage but also would open up an opportunity for better preservation of future antimicrobials in an era when their efficacy is gravely compromised and few new ones

VOL. 24, 2011

FOOD ANIMALS AND ANTIMICROBIALS: HUMAN HEALTH IMPACTS

are in the pipeline. Although the topic has been debated for several decades without definitive action, the FDA has recently made some strides in this direction. Officially, the organization now supports the conclusion that the use of medically important antimicrobials for nontherapeutic use in food animal production does not protect and promote public health (131). Although not binding, a guidance document was released in 2010 that recommended phasing in measures that would limit use of these drugs in animals and ultimately help to reduce the selection pressures that generate antimicrobial resistance (66). The Danish experience demonstrated that any negative disease effects resulting from the ban of NTAs were short-lived and that altering animal husbandry practices could counter expected increases in disease frequency (6). For aquaculture, also, it has been demonstrated that alternative processes in industry management can be instituted that will reduce antibiotic use without detrimental financial effects (141). Still, it has been argued by some in animal husbandry that the different situation in the United States will result in increased morbidity and mortality, projected to cost $1 billion or more over 10 years. Again, however, the Danish postban evaluation found that costs of production increased by just 1% for swine and were largely negligible for poultry production due to the money saved on antibiotics themselves. Models also showed that Danish swine production decreased by just 1.4% (1.7% for exports), and poultry production actually increased, by 0.4% (0.5% for exports) (158). Such calculations still fail to consider the negative externalities that are added by the burden of antibiotic resistance and the antibiotic residue pollution generated by concentrated animal feeding operations. Opponents of restriction of NTA use argue that a comprehensive risk assessment is lacking, but such an analysis is impossible without the kind of data that would come out of surveillance systems. Although surveillance systems have been advocated repeatedly (23, 70), such systems are sparse and extremely limited in their scope. In 2002, working with the accumulated evidence and an assessment of knowledge deficits in the area of animal antibiotic use, the APUA developed a set of guidelines that are still viable today and can be used to guide both policy and research agendas. In summary, APUA recommended that antimicrobials should be used only in the presence of disease, and only when prescribed by a veterinarian; that quantitative data on antimicrobial use in agriculture should be made available; that the ecology of antimicrobial resistance in agriculture should be a research priority and should be considered by regulatory agencies in assessing associated human health risks; and that efforts should be invested in improving and expanding surveillance programs for antimicrobial resistance. Suitable alternatives to NTAs can be implemented, such as vaccination, alterations in herd management, and other changes, such as targeted use of antimicrobials with a more limited dosage and duration so as not to select for resistance to critical human therapeutics (23). There is no doubt that human misuse and overuse of antibiotics are large contributors to resistance, particularly in relation to bacteria associated with human infection. Interventions in medical settings and the community are clearly needed to preserve the efficacy of antibiotics. Efforts in this area are being pursued by the Centers for Disease Control and Preven-

729

tion, the Alliance for the Prudent Use of Antibiotics, the American Medical Association, the American Academy of Pediatrics, the Infectious Diseases Society of America, and other professional groups. Still, given the large quantity of antibiotics used in food animals for nontherapeutic reasons, some measure of control over a large segment of antibiotic use and misuse can be gained by establishing guidelines for animals that permit therapeutic use only and by then tracking use and health outcomes. The current science provides overwhelming evidence that antibiotic use is a powerful selector of resistance that can appear not only at the point of origin but also nearly everywhere else (104). The latter phenomenon occurs because of the enormous ramifications of horizontal gene transfer. A mounting body of evidence shows that antimicrobial use in animals, including the nontherapeutic use of antimicrobials, leads to the propagation and shedding of substantial amounts of antimicrobial-resistant bacteria—both as pathogens, which can directly and indirectly infect humans, and as commensals, which may carry transferable resistance determinants across species borders and reach humans through multiple routes of transfer. These pathways include not only food but also water and sludge and manure applications to food crop soils. Continued nontherapeutic use of antimicrobials in food animals will increase the pool of resistance genes, as well as their density, as bacteria migrate into the environment at large. The lack of species barriers for gene transmission argues that the focus of research efforts should be directed toward the genetic infrastructure and that it is now imperative to take an ecological approach toward addressing the impacts of NTA use on human disease. The study of animal-to-human transmission of antibiotic resistance therefore requires a greater understanding of the genetic interaction and spread that occur in the larger arena of commensal and environmental bacteria. ACKNOWLEDGMENTS B. M. Marshall was supported in part by The Pew Charitable Trusts. S. B. Levy is a consultant. We thank Amadea Britton for research help. REFERENCES 1. Aarestrup, F. M. 1999. Association between the consumption of antimicrobial agents in animal husbandry and the occurrence of resistant bacteria among food animals. Int. J. Antimicrob. Agents 12:279–285. 2. Aarestrup, F. M., Y. Agerso, P. Gerner-Smidt, M. Madsen, and L. B. Jensen. 2000. Comparison of antimicrobial resistance phenotypes and resistance genes in Enterococcus faecalis and Enterococcus faecium from humans in the community, broilers, and pigs in Denmark. Diagn. Microbiol. Infect. Dis. 37:127–137. 3. Aarestrup, F. M., et al. 1996. Glycopeptide susceptibility among Danish Enterococcus faecium and Enterococcus faecalis isolates of animal and human origin and PCR identification of genes within the VanA cluster. Antimicrob. Agents Chemother. 40:1938–1940. 4. Aarestrup, F. M., F. Bager, and J. S. Andersen. 2000. Association between the use of avilamycin for growth promotion and the occurrence of resistance among Enterococcus faecium from broilers: epidemiological study and changes over time. Microb. Drug Resist. 6:71–75. 5. Aarestrup, F. M., and B. Carstensen. 1998. Effect of tylosin used as a growth promoter on the occurrence of macrolide-resistant enterococci and staphylococci in pigs. Microb. Drug Resist. 4:307–312. 6. Aarestrup, F. M., V. F. Jensen, H. D. Emborg, E. Jacobsen, and H. C. Wegener. 2010. Changes in the use of antimicrobials and the effects on productivity of swine farms in Denmark. Am. J. Vet. Res. 71:726–733. 7. Aarestrup, F. M., et al. 2001. Effect of abolishment of the use of antimicrobial agents for growth promotion on occurrence of antimicrobial resistance in fecal enterococci from food animals in Denmark. Antimicrob. Agents Chemother. 45:2054–2059.

730

MARSHALL AND LEVY

8. Akwar, T. H., et al. 2007. Risk factors for antimicrobial resistance among fecal Escherichia coli from residents on forty-three swine farms. Microb. Drug Resist. 13:69–76. 9. Alexander, T. W., et al. 2010. Farm-to-fork characterization of Escherichia coli associated with feedlot cattle with a known history of antimicrobial use. Int. J. Food Microbiol. 137:40–48. 10. Alexander, T. W., et al. 2008. Effect of subtherapeutic administration of antibiotics on the prevalence of antibiotic-resistant Escherichia coli bacteria in feedlot cattle. Appl. Environ. Microbiol. 74:4405–4416. 11. Alpharma, Inc., Animal Health. 2007. Straight talk about antibiotic use in food-producing animals, p. 1–4. In For the record, vol. 6. Alpharma, Inc., Animal Health, Bridgewater, NJ. http://antibiotictruths.com/FTR/FTR _aug07.pdf. 12. Anderson, S. A., R. W. Yeaton Woo, and L. M. Crawford. 2001. Risk assessment of the impact on human health of resistant Campylobacter jejuni from fluoroquinolone use in beef cattle. Food Control 12:13–25. 13. Angulo, F. J., V. N. Nargund, and T. C. Chiller. 2004. Evidence of an association between use of anti-microbial agents in food animals and antimicrobial resistance among bacteria isolated from humans and the human health consequences of such resistance. J. Vet. Med. B Infect. Dis. Vet. Public Health 51:374–379. 14. Anonymous. 2010. Hearing on antibiotic resistance and the use of antibiotics in animal agriculture. House Energy and Commerce Committee, Washington, DC. http://energycommerce.house.gov/hearings/hearingdetail .aspx?NewsID⫽8001. 15. Arnold, S., B. Gassner, T. Giger, and R. Zwahlen. 2004. Banning antimicrobial growth promoters in feedstuffs does not result in increased therapeutic use of antibiotics in medicated feed in pig farming. Pharmacoepidemiol. Drug Saf. 13:323–331. 16. Aubry-Damon, H., et al. 2004. Antimicrobial resistance in commensal flora of pig farmers. Emerg. Infect. Dis. 10:873–879. 17. Bager, F., F. M. Aarestrup, M. Madsen, and H. C. Wegener. 1999. Glycopeptide resistance in Enterococcus faecium from broilers and pigs following discontinued use of avoparcin. Microb. Drug Resist. 5:53–56. 18. Bager, F., M. Madsen, J. Christensen, and F. M. Aarestrup. 1997. Avoparcin used as a growth promoter is associated with the occurrence of vancomycin-resistant Enterococcus faecium on Danish poultry and pig farms. Prev. Vet. Med. 31:95–112. 19. Bailar, J. C., III, and K. Travers. 2002. Review of assessments of the human health risk associated with the use of antimicrobial agents in agriculture. Clin. Infect. Dis. 34(Suppl. 3):S135–S143. 20. Bartholomew, M. J., D. J. Vose, L. R. Tollefson, and C. C. Travis. 2005. A linear model for managing the risk of antimicrobial resistance originating in food animals. Risk Anal. 25:99–108. 21. Bartlett, J. 15 December 2003. Antibiotics and livestock: control of VRE in Denmark. Medscape Infect. Dis. http://www.medscape.com/viewarticle /465032?rss. 22. Barza, M. D., et al. (ed.). 2002. The need to improve antimicrobial use in agriculture: ecological and human health consequences. Clinical infectious diseases, vol. 34, suppl. 3. Infectious Diseases Society of America, Arlington, VA. 23. Barza, M. D., et al. (ed.). 2002. Policy recommendations. Clin. Infect. Dis. 34:S76–S77. 24. Bates, J., J. Z. Jordens, and D. T. Griffiths. 1994. Farm animals as a putative reservoir for vancomycin-resistant enterococcal infection in man. J. Antimicrob. Chemother. 34:507–514. 25. Benbrook, C. M. 2002. Antibiotic drug use in U.S. aquaculture. The Northwest Science and Environmental Policy Center, Sandpoint, ID. http://www .healthobservatory.org/library.cfm?RefID⫽37397. 26. Berge, A. C., E. R. Atwill, and W. M. Sischo. 2005. Animal and farm influences on the dynamics of antibiotic resistance in faecal Escherichia coli in young dairy calves. Prev. Vet. Med. 69:25–38. 27. Berge, A. C., D. A. Moore, T. E. Besser, and W. M. Sischo. 2009. Targeting therapy to minimize antimicrobial use in preweaned calves: effects on health, growth, and treatment costs. J. Dairy Sci. 92:4707–4714. 28. Bertrand, S., et al. 2006. Clonal emergence of extended-spectrum betalactamase (CTX-M-2)-producing Salmonella enterica serovar Virchow isolates with reduced susceptibilities to ciprofloxacin among poultry and humans in Belgium and France (2000 to 2003). J. Clin. Microbiol. 44:2897– 2903. 29. Boerlin, P., A. Wissing, F. M. Aarestrup, J. Frey, and J. Nicolet. 2001. Antimicrobial growth promoter ban and resistance to macrolides and vancomycin in enterococci from pigs. J. Clin. Microbiol. 39:4193–4195. 30. Borgen, K., M. Sorum, H. Kruse, and Y. Wasteson. 2000. Persistence of vancomycin-resistant enterococci (VRE) on Norwegian broiler farms. FEMS Microbiol. Lett. 191:255–258. 31. Brooks, J. P., and M. R. McLaughlin. 2009. Antibiotic resistant bacterial profiles of anaerobic swine lagoon effluent. J. Environ. Qual. 38:2431–2437. 32. Butaye, P., L. A. Devriese, and F. Haesebrouck. 2003. Antimicrobial growth promoters used in animal feed: effects of less well known antibiotics on gram-positive bacteria. Clin. Microbiol. Rev. 16:175–188. 33. Bywater, R., et al. 2004. A European survey of antimicrobial susceptibility

CLIN. MICROBIOL. REV. among zoonotic and commensal bacteria isolated from food-producing animals. J. Antimicrob. Chemother. 54:744–754. 34. Bywater, R. J. 2005. Identification and surveillance of antimicrobial resistance dissemination in animal production. Poult. Sci. 84:644–648. 35. Cabello, F. C. 2004. Antibiotics and aquaculture in Chile: implications for human and animal health. Rev. Med. Chil. 132:1001–1006. 36. Cabello, F. C. 2006. Heavy use of prophylactic antibiotics in aquaculture: a growing problem for human and animal health and for the environment. Environ. Microbiol. 8:1137–1144. 37. Campagnolo, E. R., et al. 2002. Antimicrobial residues in animal waste and water resources proximal to large-scale swine and poultry feeding operations. Sci. Total Environ. 299:89–95. 38. Carnevale, R. A. 2005. Antimicrobial use in food animals and human health. Med. Mal. Infect. 35:105–106. 39. Castanon, J. I. 2007. History of the use of antibiotic as growth promoters in European poultry feeds. Poult. Sci. 86:2466–2471. 40. CDC. 2010. Get smart: know when antibiotics work. Centers for Disease Control, Atlanta, GA. www.cdc.gov/Features/GetSmart. 41. Chapin, A., A. Rule, K. Gibson, T. Buckley, and K. Schwab. 2005. Airborne multidrug-resistant bacteria isolated from a concentrated swine feeding operation. Environ. Health Perspect. 113:137–142. 42. Chaslus-Dancla, E., P. Pohl, M. Meurisse, M. Marin, and J. P. Lafont. 1991. High genetic homology between plasmids of human and animal origins conferring resistance to the aminoglycosides gentamicin and apramycin. Antimicrob. Agents Chemother. 35:590–593. 43. Chee-Sanford, J. C., R. I. Aminov, I. J. Krapac, N. Garrigues-Jeanjean, and R. I. Mackie. 2001. Occurrence and diversity of tetracycline resistance genes in lagoons and groundwater underlying two swine production facilities. Appl. Environ. Microbiol. 67:1494–1502. 44. Chiller, T. M., T. Barrett, and F. J. Angulo. 2004. CDC studies incorrectly summarized in ‘critical review.’ J. Antimicrob. Chemother. 54:275–276. 45. Costa, P. M., A. Bica, P. Vaz-Pires, and F. Bernardo. 2010. Changes in antimicrobial resistance among faecal enterococci isolated from growing broilers prophylactically medicated with three commercial antimicrobials. Prev. Vet. Med. 93:71–76. 46. Cox, L. A., Jr., and D. A. Popken. 2010. Assessing potential human health hazards and benefits from subtherapeutic antibiotics in the United States: tetracyclines as a case study. Risk Anal. 30:432–457. 47. Cox, L. A., Jr., and D. A. Popken. 2004. Quantifying human health risks from virginiamycin used in chickens. Risk Anal. 24:271–288. 48. Cox, L. A., Jr., and P. F. Ricci. 2008. Causal regulations vs. political will: why human zoonotic infections increase despite precautionary bans on animal antibiotics. Environ. Int. 34:459–475. 49. DANMAP. 2008. Consumption of antimicrobial agents and the occurrence of antimicrobial resistance in bacteria from food animals, foods and humans in Denmark. DANMAP, Denmark. http://www.danmap.org/pdfFiles /Danmap_2008.pdf. 50. de Boer, E., et al. 2009. Prevalence of methicillin-resistant Staphylococcus aureus in meat. Int. J. Food Microbiol. 134:52–56. 51. de Neeling, A. J., et al. 2007. High prevalence of methicillin resistant Staphylococcus aureus in pigs. Vet. Microbiol. 122:366–372. 52. Diarra, M. S., et al. 2007. Impact of feed supplementation with antimicrobial agents on growth performance of broiler chickens, Clostridium perfringens and enterococcus counts, and antibiotic resistance phenotypes and distribution of antimicrobial resistance determinants in Escherichia coli isolates. Appl. Environ. Microbiol. 73:6566–6576. 53. Dibner, J. J., and J. D. Richards. 2005. Antibiotic growth promoters in agriculture: history and mode of action. Poult. Sci. 84:634–643. 54. Dolliver, H., K. Kumar, and S. Gupta. 2007. Sulfamethazine uptake by plants from manure-amended soil. J. Environ. Qual. 36:1224–1230. 55. Donabedian, S. M., et al. 2003. Molecular characterization of gentamicinresistant enterococci in the United States: evidence of spread from animals to humans through food. J. Clin. Microbiol. 41:1109–1113. 55a.Dumonceaux, T. J., J. E. Hill, S. M. Hemmingsen, and A. G. Van Kessel. 2006. Characterization of intestinal microbiota and response to dietary virginiamycin supplementation in the broiler chicken. Appl. Environ. Microbiol. 72:2815–2833. 56. Duran, G. M., and D. L. Marshall. 2005. Ready-to-eat shrimp as an international vehicle of antibiotic-resistant bacteria. J. Food Prot. 68:2395–2401. 57. Dutil, L., et al. 2010. Ceftiofur resistance in Salmonella enterica serovar Heidelberg from chicken meat and humans, Canada. Emerg. Infect. Dis. 16:48–54. 58. Ekkelenkamp, M. B., M. Sekkat, N. Carpaij, A. Troelstra, and M. J. Bonten. 2006. Endocarditis due to meticillin-resistant Staphylococcus aureus originating from pigs. Ned. Tijdschr. Geneeskd. 150:2442–2447. 59. Emborg, H. D., et al. 2003. Relations between the occurrence of resistance to antimicrobial growth promoters among Enterococcus faecium isolated from broilers and broiler meat. Int. J. Food Microbiol. 84:273–284. 60. Emborg, H. D., et al. 2007. Tetracycline consumption and occurrence of tetracycline resistance in Salmonella typhimurium phage types from Danish pigs. Microb. Drug Resist. 13:289–294. 61. Endtz, H. P., et al. 1991. Quinolone resistance in campylobacter isolated

VOL. 24, 2011

62.

63.

64.

65.

66.

67.

68. 69. 70. 71. 72.

73. 74. 75.

76.

77.

78.

79. 80.

81.

82. 83. 84. 85.

86.

87.

88.

89.

FOOD ANIMALS AND ANTIMICROBIALS: HUMAN HEALTH IMPACTS

from man and poultry following the introduction of fluoroquinolones in veterinary medicine. J. Antimicrob. Chemother. 27:199–208. Ervik, A., B. Thorsen, V. Eriksen, B. T. Lunestad, and O. B. Samuelsen. 1994. Impact of administering antibacterial agents on wild fish and blue mussels Mytilus edulis in the vicinity of fish farms. Dis. Aquat. Org. 18:45– 51. Fairchild, A. S., et al. 2005. Effects of orally administered tetracycline on the intestinal community structure of chickens and on tet determinant carriage by commensal bacteria and Campylobacter jejuni. Appl. Environ. Microbiol. 71:5865–5872. FAO. 2007. The state of world fisheries and aquaculture 2006. Fisheries and Aquaculture Department, Food and Agriculture Organisation of the United Nations, Rome, Italy. http://www.fao.org/docrep/009/a0699e /A0699E00.htm. FDA. 2000. FDA Task Force on Antimicrobial Resistance: key recommendations and report, Washington, DC. FDA, Washington, DC. http://www .fda.gov/downloads/ForConsumers/ConsumerUpdates/UCM143458.pdf. FDA. 2010. The judicious use of medically important antimicrobial drugs in food-producing animals. Draft guidance 209. Center for Veterinary Medicine, FDA, Washington, DC. http://www.fda.gov/downloads/AnimalVeterinary /GuidanceComplianceEnforcement/GuidanceforIndustry/UCM216936.pdf. FDA. 2009. Summary report on antimicrobials sold or distributed for use in food-producing animals. www.fda.gov/downloads/ForIndustry/UserFees /AnimalDrugUserFeeActADUFA/UCM231851.pdf. Ferber, D. 2002. Livestock feed ban preserves drug’s power. Science 295: 27–28. Fey, P. D., et al. 2000. Ceftriaxone-resistant salmonella infection acquired by a child from cattle. N. Engl. J. Med. 342:1242–1249. Franklin, A. 1999. Current status of antibiotic resistance in animal production. Acta Vet. Scand. Suppl. 92:23–28. Fuhrman, J. A. 1999. Marine viruses and their biogeochemical and ecological effects. Nature 399:541–548. Gibbs, S. G., et al. 2006. Isolation of antibiotic-resistant bacteria from the air plume downwind of a swine confined or concentrated animal feeding operation. Environ. Health Perspect. 114:1032–1037. Gilbert, J. M., D. G. White, and P. F. McDermott. 2007. The US national antimicrobial resistance monitoring system. Future Microbiol. 2:493–500. Gorbach, S. L. 2001. Antimicrobial use in animal feed—time to stop. N. Engl. J. Med. 345:1202–1203. Graslund, S., K. Holmstrom, and A. Wahlstrom. 2003. A field survey of chemicals and biological products used in shrimp farming. Mar. Pollut. Bull. 46:81–90. Grave, K., V. F. Jensen, K. Odensvik, M. Wierup, and M. Bangen. 2006. Usage of veterinary therapeutic antimicrobials in Denmark, Norway and Sweden following termination of antimicrobial growth promoter use. Prev. Vet. Med. 75:123–132. Grave, K., M. C. Kaldhusdal, H. Kruse, L. M. Harr, and K. Flatlandsmo. 2004. What has happened in Norway after the ban of avoparcin? Consumption of antimicrobials by poultry. Prev. Vet. Med. 62:59–72. Graveland, H., et al. 2010. Methicillin resistant Staphylococcus aureus ST398 in veal calf farming: human MRSA carriage related with animal antimicrobial usage and farm hygiene. PLoS One 5:e10990. Gupta, A., et al. 2004. Antimicrobial resistance among Campylobacter strains, United States, 1997–2001. Emerg. Infect. Dis. 10:1102–1109. Hammerum, A. M., V. Fussing, F. M. Aarestrup, and H. C. Wegener. 2000. Characterization of vancomycin-resistant and vancomycin-susceptible Enterococcus faecium isolates from humans, chickens and pigs by RiboPrinting and pulsed-field gel electrophoresis. J. Antimicrob. Chemother. 45:677– 680. Hamscher, G., H. T. Pawelzick, S. Sczesny, H. Nau, and J. Hartung. 2003. Antibiotics in dust originating from a pig-fattening farm: a new source of health hazard for farmers? Environ. Health Perspect. 111:1590–1594. Hanselman, B. A., et al. 2006. Methicillin-resistant Staphylococcus aureus colonization in veterinary personnel. Emerg. Infect. Dis. 12:1933–1938. Hershberger, E., et al. 2005. Epidemiology of antimicrobial resistance in enterococci of animal origin. J. Antimicrob. Chemother. 55:127–130. Heuer, O. E., et al. 2009. Human health consequences of use of antimicrobial agents in aquaculture. Clin. Infect. Dis. 49:1248–1253. Heuer, O. E., K. Pedersen, L. B. Jensen, M. Madsen, and J. E. Olsen. 2002. Persistence of vancomycin-resistant enterococci (VRE) in broiler houses after the avoparcin ban. Microb. Drug Resist. 8:355–361. Ho, P. L., et al. 2010. Genetic identity of aminoglycoside-resistance genes in Escherichia coli isolates from human and animal sources. J. Med. Microbiol. 59:702–707. Holmberg, S. D., M. T. Osterholm, K. A. Senger, and M. L. Cohen. 1984. Drug-resistant Salmonella from animals fed antimicrobials. N. Engl. J. Med. 311:617–622. Holmberg, S. D., J. G. Wells, and M. L. Cohen. 1984. Animal-to-man transmission of antimicrobial-resistant Salmonella: investigations of U.S. outbreaks, 1971–1983. Science 225:833–835. Howells, C. H., and D. H. Joynson. 1975. Possible role of animal feedingstuffs in spread of antibiotic-resistant intestinal coliforms. Lancet i:156–157.

731

90. Hummel, R., H. Tschape, and W. Witte. 1986. Spread of plasmid-mediated nourseothricin resistance due to antibiotic use in animal husbandry. J. Basic Microbiol. 26:461–466. 91. Hurd, H. S., and S. Malladi. 2008. A stochastic assessment of the public health risks of the use of macrolide antibiotics in food animals. Risk Anal. 28:695–710. 92. Inglis, G. D., et al. 2005. Effects of subtherapeutic administration of antimicrobial agents to beef cattle on the prevalence of antimicrobial resistance in Campylobacter jejuni and Campylobacter hyointestinalis. Appl. Environ. Microbiol. 71:3872–3881. 93. Jackson, C. R., P. J. Fedorka-Cray, J. B. Barrett, and S. R. Ladely. 2004. Effects of tylosin use on erythromycin resistance in enterococci isolated from swine. Appl. Environ. Microbiol. 70:4205–4210. 94. Johnson, J. R., et al. 2006. Similarity between human and chicken Escherichia coli isolates in relation to ciprofloxacin resistance status. J. Infect. Dis. 194:71–78. 95. Johnson, J. R., et al. 2007. Antimicrobial drug-resistant Escherichia coli from humans and poultry products, Minnesota and Wisconsin, 2002–2004. Emerg. Infect. Dis. 13:838–846. 96. Jones, F. T., and S. C. Ricke. 2003. Observations on the history of the development of antimicrobials and their use in poultry feeds. Poult. Sci. 82:613–617. 97. Kaszanyitzky, E. J., M. Tenk, A. Ghidan, G. Y. Fehervari, and M. Papp. 2007. Antimicrobial susceptibility of enterococci strains isolated from slaughter animals on the data of Hungarian resistance monitoring system from 2001 to 2004. Int. J. Food Microbiol. 115:119–123. 98. Katsunuma, Y., et al. 2007. Associations between the use of antimicrobial agents for growth promotion and the occurrence of antimicrobial-resistant Escherichia coli and enterococci in the feces of livestock and livestock farmers in Japan. J. Gen. Appl. Microbiol. 53:273–279. 99. Kelly, L., et al. 2004. Animal growth promoters: to ban or not to ban? A risk assessment approach. Int. J. Antimicrob. Agents 24:205–212. 100. Khachatryan, A. R., T. E. Besser, D. D. Hancock, and D. R. Call. 2006. Use of a nonmedicated dietary supplement correlates with increased prevalence of streptomycin-sulfa-tetracycline-resistant Escherichia coli on a dairy farm. Appl. Environ. Microbiol. 72:4583–4588. 101. Khanna, T., R. Friendship, C. Dewey, and J. S. Weese. 2008. Methicillin resistant Staphylococcus aureus colonization in pigs and pig farmers. Vet. Microbiol. 128:298–303. 102. Klare, I., et al. 1999. Decreased incidence of VanA-type vancomycin-resistant enterococci isolated from poultry meat and from fecal samples of humans in the community after discontinuation of avoparcin usage in animal husbandry. Microb. Drug Resist. 5:45–52. 103. Kriebel, D., et al. 2001. The precautionary principle in environmental science. Environ. Health Perspect. 109:871–876. 104. Kruse, H., and H. Sorum. 1994. Transfer of multiple drug resistance plasmids between bacteria of diverse origins in natural microenvironments. Appl. Environ. Microbiol. 60:4015–4021. 105. Kumar, K., S. C. Gupta, Y. Chander, and A. K. Singh. 2005. Antibiotic use in agriculture and its impact on the terrestrial environment. Adv. Agron. 87:1–54. 106. Labbate, M., R. J. Case, and H. W. Stokes. 2009. The integron/gene cassette system: an active player in bacterial adaptation. Methods Mol. Biol. 532: 103–125. 107. Lauderdale, T. L., et al. 2007. Effect of banning vancomycin analogue avoparcin on vancomycin-resistant enterococci in chicken farms in Taiwan. Environ. Microbiol. 9:819–823. 108. Lee, J. H. 2003. Methicillin (oxacillin)-resistant Staphylococcus aureus strains isolated from major food animals and their potential transmission to humans. Appl. Environ. Microbiol. 69:6489–6494. 109. Levy, S. B. 2002. The antibiotic paradox: how the misuse of antibiotics destroys their curative powers, 2nd ed. Perseus Publishing, Cambridge, MA. 110. Levy, S. B. 1998. The challenge of antibiotic resistance. Sci. Am. 278:46–53. 111. Levy, S. B., G. B. FitzGerald, and A. B. Macone. 1976. Changes in intestinal flora of farm personnel after introduction of a tetracycline-supplemented feed on a farm. N. Engl. J. Med. 295:583–588. 112. Levy, S. B., G. B. FitzGerald, and A. B. Macone. 1976. Spread of antibioticresistant plasmids from chicken to chicken and from chicken to man. Nature 260:40–42. 113. Levy, S. B., and B. Marshall. 2004. Antibacterial resistance worldwide: causes, challenges and responses. Nat. Med. 10:S122–S129. 114. Lewis, H. C., et al. 2008. Pigs as source of methicillin-resistant Staphylococcus aureus CC398 infections in humans, Denmark. Emerg. Infect. Dis. 14:1383–1389. 115. Luangtongkum, T., et al. 2006. Effect of conventional and organic production practices on the prevalence and antimicrobial resistance of Campylobacter spp. in poultry. Appl. Environ. Microbiol. 72:3600–3607. 116. Marshall, B. M., D. J. Ochieng, and S. B. Levy. 2009. Commensals: underappreciated reservoirs of resistance. Microbe 4:231–238. 117. Mellon, M., C. Benbrook, and K. L. Benbrook. 2001. Hogging it: estimates of antimicrobial abuse in livestock. USC Publications, Cambridge, MA. 118. Molbak, K., et al. 1999. An outbreak of multidrug-resistant, quinolone-

732

119.

120.

121.

122.

123.

124.

125.

126.

127.

128.

129.

130.

131.

132.

133. 134.

135.

136.

137. 138.

139.

140. 141.

142.

MARSHALL AND LEVY resistant Salmonella enterica serotype Typhimurium DT104. N. Engl. J. Med. 341:1420–1425. Noble, W. C., Z. Virani, and R. G. Cree. 1992. Co-transfer of vancomycin and other resistance genes from Enterococcus faecalis NCTC 12201 to Staphylococcus aureus. FEMS Microbiol. Lett. 72:195–198. Normanno, G., et al. 2007. Methicillin-resistant Staphylococcus aureus (MRSA) in foods of animal origin product in Italy. Int. J. Food Microbiol. 117:219–222. Pantosti, A., M. Del Grosso, S. Tagliabue, A. Macri, and A. Caprioli. 1999. Decrease of vancomycin-resistant enterococci in poultry meat after avoparcin ban. Lancet 354:741–742. Perreten, V. 2005. Resistance in the food chain and in bacteria from animals: relevance to human infections, p. 446–464. In D. G. White, M. N. Alekshun, and P. F. McDermott (ed.), Frontiers in antimicrobial resistance: a tribute to Stuart B. Levy. ASM Press, Washington, DC. Petersen, A., J. S. Andersen, T. Kaewmak, T. Somsiri, and A. Dalsgaard. 2002. Impact of integrated fish farming on antimicrobial resistance in a pond environment. Appl. Environ. Microbiol. 68:6036–6042. Phillips, I., et al. 2004. Does the use of antibiotics in food animals pose a risk to human health? A critical review of published data. J. Antimicrob. Chemother. 53:28–52. Post, V., G. D. Recchia, and R. M. Hall. 2007. Detection of gene cassettes in Tn402-like class 1 integrons. Antimicrob. Agents Chemother. 51:3467– 3468. Price, L. B., et al. 2007. Elevated risk of carrying gentamicin-resistant Escherichia coli among U.S. poultry workers. Environ. Health Perspect. 115:1738–1742. Rhodes, G., et al. 2000. Distribution of oxytetracycline resistance plasmids between aeromonads in hospital and aquaculture environments: implication of Tn1721 in dissemination of the tetracycline resistance determinant tet(A). Appl. Environ. Microbiol. 66:3883–3890. Sanchez, R., et al. 1994. Evolution of susceptibilities of Campylobacter spp. to quinolones and macrolides. Antimicrob. Agents Chemother. 38:1879– 1882. Sapkota, A. R. 2006. Antibiotic resistance genes in drug resistant Enterococcus spp and Streptococcus spp recovered from indoor air of a concentrated swine-feeding operation. Lett. Appl. Microbiol. 43:534–540. Schroeder, C. M., et al. 2003. Isolation of antimicrobial-resistant Escherichia coli from retail meats purchased in Greater Washington, DC, U.S.A. Int. J. Food Microbiol. 85:197–202. Sharfstein, J. M. 14 July 2010. Testimony. FDA, Silver Spring, MD. http: //democrats.energycommerce.house.gov/documents/20100714/Sharfstein .Testimony.07.14.2010.pdf. Sharma, R., et al. 2008. Diversity and distribution of commensal fecal Escherichia coli bacteria in beef cattle administered selected subtherapeutic antimicrobials in a feedlot setting. Appl. Environ. Microbiol. 74:6178–6186. Shotwell, T. K. (ed.). 2009. Superbugs: E. coli, Salmonella, Staphylococcus and more!, 1st ed. Biontogeny Publications, Bridgeport, TX. Smith, D. L., A. D. Harris, J. A. Johnson, E. K. Silbergeld, and J. G. Morris, Jr. 2002. Animal antibiotic use has an early but important impact on the emergence of antibiotic resistance in human commensal bacteria. Proc. Natl. Acad. Sci. U. S. A. 99:6434–6439. Smith, D. L., et al. 2003. Assessing risks for a pre-emergent pathogen: virginiamycin use and the emergence of streptogramin resistance in Enterococcus faecium. Lancet Infect. Dis. 3:241–249. Smith, H. W., and W. E. Crabb. 1957. The effect of the continuous administration of diets containing low levels of tetracyclines on the incidence of drug-resistant Bacterium coli in the faeces of pigs and chickens: the sensitivity of the Bact. coli to other chemotherapeutic agents. Vet. Rec. 69:24– 30. Smith, K. E., et al. 1999. Quinolone-resistant Campylobacter jejuni infections in Minnesota, 1992–1998. N. Engl. J. Med. 340:1525–1532. Smith, T. C., et al. 2009. Methicillin-resistant Staphylococcus aureus (MRSA) strain ST398 is present in Midwestern U.S. swine and swine workers. PLoS One 4:e4258. Sorensen, T. L., et al. 2001. Transient intestinal carriage after ingestion of antibiotic-resistant Enterococcus faecium from chicken and pork. N. Engl. J. Med. 345:1161–1166. Sorum, H. 1999. Antibiotic resistance in aquaculture. Acta Vet. Scand. Suppl. 92:29–36. Sorum, H. 2006. Antimicrobial drug resistance in fish pathogens, p. 213– 238. In F. M. Aarestrup (ed.), Antimicrobial resistance in bacteria of animal origin. ASM Press, Washington, DC. Stokestad, E. L. R., and T. H. Jukes. 1950. Further observations on the “animal protein factor.” Proc. Soc. Exp. Biol. Med. 73:523–528.

CLIN. MICROBIOL. REV. 143. Summers, A. O. 2002. Generally overlooked fundamentals of bacterial genetics and ecology. Clin. Infect. Dis. 34(Suppl. 3):S85–S92. 144. Swann, M. 1969. Report of the Joint Committee on the Use of Antibiotics in Animal Husbandry and Veterinary Medicine. Her Majesty’s Stationery Office, London, United Kingdom. 145. Tacket, C. O., L. B. Dominguez, H. J. Fisher, and M. L. Cohen. 1985. An outbreak of multiple-drug-resistant Salmonella enteritis from raw milk. JAMA 253:2058–2060. 146. Tschape, H. 1994. The spread of plasmids as a function of bacterial adaptability. FEMS Microbiol. Ecol. 15:23–31. 147. van Belkum, A., et al. 2008. Methicillin-resistant and -susceptible Staphylococcus aureus sequence type 398 in pigs and humans. Emerg. Infect. Dis. 14:479–483. 148. van den Bogaard, A. E., N. Bruinsma, and E. E. Stobberingh. 2000. The effect of banning avoparcin on VRE carriage in The Netherlands. J. Antimicrob. Chemother. 46:146–148. 149. van den Bogaard, A. E., R. Willems, N. London, J. Top, and E. E. Stobberingh. 2002. Antibiotic resistance of faecal enterococci in poultry, poultry farmers and poultry slaughterers. J. Antimicrob. Chemother. 49:497–505. 150. van Leeuwen, W. J., P. A. Guinee, C. E. Voogd, and B. van Klingeren. 1986. Resistance to antibiotics in Salmonella. Tijdschr. Diergeneeskd. 111:9–13. 151. Van Loo, I. H. 2007. Emergence of methicillin-resistant Staphylococcus aureus of animal origins in humans. Emerg. Infect. Dis. 13:1834–1839. 152. van Rijen, M. M., P. H. Van Keulen, and J. A. Kluytmans. 2008. Increase in a Dutch hospital of methicillin-resistant Staphylococcus aureus related to animal farming. Clin. Infect. Dis. 46:261–263. 153. Varga, C., et al. 2009. Associations between reported on-farm antimicrobial use practices and observed antimicrobial resistance in generic fecal Escherichia coli isolated from Alberta finishing swine farms. Prev. Vet. Med. 88:185–192. 154. Varga, C., A. Rajic, M. E. McFall, R. J. Reid-Smith, and S. A. McEwen. 2009. Associations among antimicrobial use and antimicrobial resistance of Salmonella spp. isolates from 60 Alberta finishing swine farms. Foodborne Pathog. Dis. 6:23–31. 155. Voss, A., F. Loeffen, J. Bakker, C. Klaassen, and M. Wulf. 2005. Methicillinresistant Staphylococcus aureus in pig farming. Emerg. Infect. Dis. 11:1965– 1966. 156. Weber, J. T., et al. 1994. Epidemic cholera in Ecuador: multidrug-resistance and transmission by water and seafood. Epidemiol. Infect. 112:1–11. 156a.Wegener, H. C. 2003. Ending the use of antimicrobial growth promoters is making a difference. ASM News 69:443–448. 157. WHO. June 2006. Antimicrobial use in aquaculture and antimicrobial resistance. Report of a joint FAO/OIE/WHO expert consultation on antimicrobial use in aquaculture and antimicrobial resistance. WHO, Geneva, Switzerland. ftp://ftp.fao.org/ag/agn/food/aquaculture_rep_13_16june2006 .pdf. 158. WHO. 2002. Impacts of antimicrobial growth promoter termination in Denmark. Department of Communicable Diseases, World Health Organization, Foulum, Denmark. http://whqlibdoc.who.int/hq/2003/WHO_CDS _CPE_ZFK_2003.1.pdf. 159. WHO. October 1997. The medical impact of the use of antimicrobials in food animals: report of a WHO meeting. World Health Organization, Geneva, Switzerland. http://whqlibdoc.who.int/hq/1997/WHO_EMC_ZOO _97.4.pdf. 160. Wierup, M. 2001. The Swedish experience of the 1986 year ban of antimicrobial growth promoters, with special reference to animal health, disease prevention, productivity, and usage of antimicrobials. Microb. Drug Resist. 7:183–190. 161. Winokur, P. L., D. L. Vonstein, L. J. Hoffman, E. K. Uhlenhopp, and G. V. Doern. 2001. Evidence for transfer of CMY-2 AmpC beta-lactamase plasmids between Escherichia coli and Salmonella isolates from food animals and humans. Antimicrob. Agents Chemother. 45:2716–2722. 162. Witte, W. 2000. Selective pressure by antibiotic use in livestock. Int. J. Antimicrob. Agents 16(Suppl. 1):S19–S24. 163. Witte, W., B. Strommenger, C. Stanek, and C. Cuny. 2007. Methicillinresistant Staphylococcus aureus ST398 in humans and animals, Central Europe. Emerg. Infect. Dis. 13:255–258. 164. Zhang, X. Y., L. J. Ding, and M. Z. Fan. 2009. Resistance patterns and detection of aac(3)-IV gene in apramycin-resistant Escherichia coli isolated from farm animals and farm workers in northeastern of China. Res. Vet. Sci. 87:449–454. 165. Zhao, S., et al. 2003. Characterization of Salmonella enterica serotype Newport isolated from humans and food animals. J. Clin. Microbiol. 41:5366– 5371.

VOL. 24, 2011

FOOD ANIMALS AND ANTIMICROBIALS: HUMAN HEALTH IMPACTS

Bonnie Marshall is a Senior Research Associate in the Center for Adaptation Genetics and Drug Resistance in the Department of Microbiology and Molecular Biology at Tufts University School of Medicine in Boston, MA. After obtaining a B.A. in Microbiology at the University of New Hampshire, she did work on herpesviruses at Harvard’s New England Regional Primate Research Center and then returned to school to complete a degree in medical technology. In 1977, she joined the laboratory of Dr. Stuart Levy, from which she has published over 23 peer-reviewed publications on the ecology and epidemiology of resistance genes in human and animal clinical and commensal bacteria and environmental strains of water, soils, and plants. Ms. Marshall has also been engaged actively for 30 years with the Alliance for the Prudent Use of Antibiotics, where she is Staff Scientist and serves on the Board of Directors.

733

Stuart B. Levy is a Board-Certified Internist at Tufts Medical Center, a Professor of Molecular Biology and Microbiology and of Medicine at Tufts University School of Medicine, and Director, Center for Adaptation Genetics and Drug Resistance, also at Tufts University School of Medicine. He received his B.A. degree from Williams College and his M.D. from the University of Pennsylvania. He cofounded and remains active in both The Alliance for the Prudent Use of Antibiotics (1981) and Paratek Pharmaceuticals, Inc. (1996). More than 4 decades of studies on the molecular, genetic, and ecologic bases of drug resistance have led to over 250 peer-reviewed publications, authorship of The Antibiotic Paradox, honorary degrees in biology from Wesleyan University (1998) and Des Moines University (2001), ASM’s Hoechst-Roussel Award for esteemed research in antimicrobial chemotherapy, and ICS’s Hamao Umezawa Memorial Award. Dr. Levy is a Past President of the American Society for Microbiology and a Fellow of the American College of Physicians (ACP), the Infectious Diseases Society of America, the American Academy of Microbiology (AAM), and the American Association for the Advancement of Science. He serves on the National Science Advisory Board for Biosecurity.

CLINICAL MICROBIOLOGY REVIEWS, Oct. 2011, p. 734–754 0893-8512/11/$12.00 doi:10.1128/CMR.00015-11 Copyright © 2011, American Society for Microbiology. All Rights Reserved.

Vol. 24, No. 4

Human Metapneumovirus: Lessons Learned over the First Decade† Verena Schildgen,1 Bernadette van den Hoogen,2 Ron Fouchier,2 Ralph A. Tripp,3 Rene Alvarez,4 Catherine Manoha,5 John Williams,6 and Oliver Schildgen1* Kliniken der Stadt Ko ¨ln gGmbH, University Hospital Witten/Herdecke, Cologne, Germany1; Department of Virology, Erasmus Medical Center Rotterdam, Rotterdam, The Netherlands2; Department of Infectious Diseases, University of Georgia, Athens, Georgia 306023; United States Naval Medical Research Unit San Antonio, Department of Dental and Biomedical Research, Applied Laboratory Sciences Division, Ft. Sam Houston, Texas 782344; Laboratoire de Virologie, Centre Hospitalier Universitaire, Dijon, France5; and Vanderbilt University Medical Center, School of Medicine, Nashville, Tennessee 37232-25816 INTRODUCTION .......................................................................................................................................................734 HMPV BIOLOGY .......................................................................................................................................................734 Description of the Agent ........................................................................................................................................734 Taxonomy .............................................................................................................................................................735 HMPV Replication..................................................................................................................................................735 EPIDEMIOLOGY .......................................................................................................................................................737 Molecular Epidemiology and Virus Evolution....................................................................................................737 Clinical Epidemiology ............................................................................................................................................739 CLINICAL FEATURES OF HMPV INFECTION .................................................................................................739 Symptoms and Pathology.......................................................................................................................................739 Risk Groups Other than Children .......................................................................................................................741 Immunosuppressed individuals ........................................................................................................................741 Older adults.........................................................................................................................................................741 Coinfections .............................................................................................................................................................742 Treatment Options .................................................................................................................................................742 IMMUNE RESPONSES IN HUMANS AND ANIMALS ......................................................................................743 Innate Immunity .....................................................................................................................................................743 Humoral Response..................................................................................................................................................743 Cellular Immunity ..................................................................................................................................................744 ANIMAL MODELS OF HMPV INFECTION ........................................................................................................744 Mouse Model ...........................................................................................................................................................744 Cotton Rat Model ...................................................................................................................................................745 Hamster and Ferret Models..................................................................................................................................745 HMPV Infection in Nonhuman Primates............................................................................................................745 VACCINE DEVELOPMENT.....................................................................................................................................745 DIAGNOSTICS ...........................................................................................................................................................746 Molecular Diagnostics............................................................................................................................................746 Fluorescent-Antibody Staining and ELISA .........................................................................................................747 Cell Culture .............................................................................................................................................................747 FUTURE PERSPECTIVES........................................................................................................................................747 ACKNOWLEDGMENTS ...........................................................................................................................................747 REFERENCES ............................................................................................................................................................747 and lower respiratory tract illnesses. In 2001, a previously unknown virus, human metapneumovirus (HMPV), was added to this list. In this review, the current knowledge on HMPV is summarized.

INTRODUCTION Respiratory tract infections (RTIs) are a leading cause of morbidity and mortality worldwide. For children under the age of 5 years old, RTIs are ranked as the second leading cause of death, regardless of geographical location (172). In children, respiratory syncytial virus (RSV), parainfluenza viruses (PIVs), and influenza virus are known major causes of bronchiolitis

HMPV BIOLOGY Description of the Agent Human metapneumovirus was first detected upon inoculation of tertiary monkey kidney cells with respiratory specimens collected from children with RTIs for which the etiological agent could not be identified using diagnostic assays for known respiratory viruses. Cytopathic effects morphologically indistinguishable from those induced by RSV were observed. Virus-

* Corresponding author. Mailing address: Dipl.-Biologe, Kliniken der Stadt Ko ¨ ln gGmbH, Krankenhaus Merheim, Klinikum der Privaten Universita¨t Witten/Herdecke, Institut fu ¨r Pathologie, Ostmerheimer Str. 200, D-51109 Cologne, Germany. Phone: 49-(0)221-890713467. Fax: 49-(0)221-8907-3542. E-mail: [email protected]. † All authors contributed equally to the manuscript. 734

VOL. 24, 2011

HUMAN METAPNEUMOVIRUS

735

13,280 to 13,378 nucleotides and contains at least 8 genes and 9 open reading frames (ORFs). Beyond the genes for the proteins mentioned above, the HMPV genome, similar to the RSV genome, contains the M2 gene, from which the M2-1 and M2-2 proteins are expressed. However, distinct from RSV, the HMPV genome lacks nonstructural genes (NS1 and NS2), and the order of genes between M and L is different (in RSV, the order is SH-G-F-M2, and in HMPV, the order is F-M2-SH-G) (230, 231) (Fig. 2). Taxonomy. Human metapneumovirus belongs to the order Mononegavirales, in the family Paramyxoviridae. HMPV was classified as the first human member of the Metapneumovirus genus, in the subfamily Pneumovirinae of the family Paramyxoviridae (231). HMPV Replication FIG. 1. Electron micrograph of HMPV particles. Virions concentrated from infected cell culture supernatants were visualized by negative-contrast electron microscopy after phosphotungstic acid staining. Magnification, ⫻92,000.

infected cell supernatants revealed pleomorphic particles measuring 150 to 600 nm, with short envelope projections of 13 to 17 nm, by electron microscopy (Fig. 1). These supernatants did not display hemagglutinating activity, and virus propagation was found to be dependent on trypsin (231). PCR and sequence analysis revealed a viral genome with close resemblance to that of avian metapneumovirus (AMPV), a virus causing swollen head syndrome and rhinotracheitis in chickens and turkeys (37). AMPVs have been classified into four subgroups, subgroups A through D (18), among which AMPV-C is most closely related to HMPV. However, the highly variable attachment (G) protein and small hydrophobic (SH) protein of AMPV and HMPV share only 20 to 30% amino acid sequence identity, while the percent sequence identity is ⬃80% for the other structural proteins (230). Similar to other pneumoviruses, HMPV virions contain a lipid membrane envelope surrounding the matrix (M) protein and three transmembrane surface glycoproteins, the fusion (F), G, and SH proteins. Within the envelope lies a helical ribonucleoprotein (RNP) complex, which consists of nucleoprotein (N), phosphoprotein (P), large polymerase protein (L), and the nonsegmented single-stranded negative-sense RNA genome. The genome size of HMPV ranges in length from

The HMPV replication cycle begins with attachment of the virus to the host cell, which is thought to be directed by the G protein (136). The G protein is the most variable protein among HMPV isolates (236). The deduced amino acid sequence of the G protein contains a single hydrophobic region that is located near the N terminus and is thought to serve as both an uncleaved signal peptide and a membrane anchor. The C-terminal three-fourths of the molecule are thought to be extracellular. The HMPV G protein has a high content of serine and threonine residues, which are potential acceptor sites for O-linked glycosylation, and a high content of proline residues (230)—features shared with heavily glycosylated mucin-like structures. The predicted structural features of the G protein were confirmed by analyses of the biosynthesis, glycosylation, intracellular transport, and cell surface expression of the G protein (144). It has been suggested that cellular glycosaminoglycans, including heparin sulfate-like molecules, are involved in the binding of the G protein to the host cell (224). Recombinant viruses lacking the G protein are able to replicate both in vitro and in vivo, indicating that attachment via the G protein is not required for subsequent steps in the replication cycle (22, 25). Fusion of the viral membrane with the host cell membrane is mediated by the F protein. The structural organization of the HMPV F protein is similar to that of other viral class I fusion proteins, where the F protein is synthesized as an F0 precursor protein that requires cleavage by proteases to yield the activated disulfide-linked F1 and F2 subunits (reviewed in reference 135). Although the cleavage motif of the HMPV F pro-

FIG. 2. Genomic maps of HMPV and RSV showing the important differences between the two viruses. Compared to HMPV, RSV expresses two extra proteins, NS1 and NS2, the positions of the SH and G proteins differ, and the reading frames for M2 and L overlap in RSV. The double diagonal lines crossing the L ORF indicate the shortened representation of the L gene. Le, leader; N, nucleoprotein; P, phosphoprotein; M, matrix protein; F, fusion protein; SH, small hydrophobic protein; G, attachment protein; L, large polymerase protein; Tr, trailer; NS1 and NS2, nonstructural proteins 1 and 2.

736

SCHILDGEN ET AL.

tein (RQSR2F) contains a minimal furin cleavage site that is typical for most paramyxoviruses (RXXR2F), the HMPV F protein appears to require exogenous protease activation, as it is not cleaved intracellularly (reviewed in reference 212). In vitro, cleavage of the HMPV F protein is facilitated by the addition of trypsin to the cell culture medium. However, some laboratory strains have been shown to replicate in the absence of exogenous trypsin, likely due to a change in the cleavage site (RQPR2F) (198), though this altered protease cleavability does not affect virulence (23). For most paramyxoviruses, fusion depends on an interaction between the F protein and its cognate attachment protein (reviewed in reference 136). However, the F proteins of members of the Pneumovirinae subfamily do not depend upon their cognate G proteins for fusion and are processed efficiently and correctly into a biologically active form when expressed in the absence of other viral proteins (114, 125, 178, 203, 220). This is consistent with the observation that HMPV lacking the G protein/gene remains replication competent in vitro and in vivo (22). Recently, it was shown that the HMPV F protein can engage in binding to host cells via integrin ␣v␤1 via a conserved Arg-Gly-Asp (RGD) motif, providing evidence for a role of the F protein in attachment in addition to membrane fusion (47). Membrane fusion promoted by paramyxovirus F proteins generally occurs at the plasma membrane of the host cell at neutral pH (134, 195), which contrasts with the case for viruses gaining entry via a pH-dependent endocytic route. Interestingly, it has been shown that syncytium formation for HMPV strain Can97-83 is promoted by the HMPV F protein at low pH, suggesting a unique mechanism of triggering fusion among the paramyxovirus F proteins (203). However, the low-pH dependency is not a general phenomenon for HMPV F proteins and appears to be restricted to a few laboratory strains that contain an E294G substitution in the F protein (114). As for most paramyxoviruses, the trigger that leads to the membrane fusion event remains unknown, although it is tempting to speculate that binding to integrin ␣v␤1 may provide such a trigger (47). By generating chimeric viruses between HMPV and AMPV, it has been shown that the F protein is an important determinant of metapneumovirus host range (57). Besides the F and G proteins, HMPV harbors a third putative transmembrane surface glycoprotein, SH. Hydrophilicity profiles of AMPV, HMPV, and RSV SH proteins were found to be similar, although the RSV SH protein appears to be truncated compared to the SH proteins of AMPV and HMPV (233). The SH protein of HMPV has a high threonine/serine content of ⬃22% and contains 10 cysteine residues (230). Recombinant HMPV lacking only the SH gene was not attenuated or showed only marginal attenuation in vitro and in animal models (12, 22, 58). Infection of mice with HMPV lacking the SH gene resulted in enhanced secretion of proinflammatory cytokines compared to that in mice infected with wild-type virus (12). However, similar analyses using human lung epithelial cells infected with wild-type HMPV or the mutant virus lacking the SH gene did not reveal differential expression of genes or proteins (58). Moreover, the latter study did not reveal any effects of the SH protein on replication kinetics, cytopathic effects, or plaque formation of HMPV. Thus, the function of the HMPV SH protein remains elusive. Following membrane fusion, the viral RNP containing the

CLIN. MICROBIOL. REV.

negative-sense viral RNA (vRNA) genome is released into the cytoplasm, where it serves as a template for the synthesis of mRNA and antigenomic cRNA. Most of the knowledge on HMPV transcription is inferred from the knowledge of RSV and other paramyxoviruses (136). The HMPV genomic termini contain leader and trailer sequences that are partially complementary and act as promoters to direct the transcription of mRNA and cRNA or vRNA, respectively. The leader and trailer of HMPV are less complementary than those of related paramyxoviruses. Their functions were confirmed in minigenome assays showing that the leader and trailer sequences were sufficient to drive replication and expression of heterologous genes when HMPV polymerase complex proteins were present (110). Using minigenome assays and recombinant virus rescue, it was further shown that the polymerase complex proteins and genomic termini of HMPV and AMPV are interchangeable, in agreement with the close genetic relationship between these two viruses (54). Nevertheless, viruses with HMPV-AMPV chimeric polymerase complexes were found to be attenuated in animal models and may thus represent useful live vaccine candidates (54, 184). Similar minigenome and reverse genetics studies further showed that while the L, P, and N proteins were absolutely required for minigenome expression or recombinant virus rescue, the M2-1 protein was dispensable (110). The M2-1 protein of RSV has been shown to promote transcription processivity (46, 78), but such a role was not observed for M2-1 of HMPV (35, 110). In addition to the M2-1 ORF, the M2 gene of pneumoviruses also contains a second ORF, the M2-2 ORF. Expression of the M2-2 protein is achieved via a process of coupled translation, a mechanism of translational initiation in which the ribosomes that translate the first ORF move a short distance upstream after termination and reinitiate translation from the second overlapping ORF (94). For RSV, the M2-2 protein is thought to play a role in shifting the balance of RNA synthesis from mRNA to vRNA (20). Recent evidence suggests that M2-2 of HMPV regulates RNA synthesis in a similar fashion (35) and potentially also affects the fidelity of the polymerase complex (197). More work is needed to gain detailed knowledge of the HMPV polymerase complexes and the roles of M2-1 and M2-2 during virus replication. The genome of HMPV further contains noncoding regions between each ORF that range in size from 23 to 209 nucleotides and contain gene end signals, intergenic regions, and gene start signals, with little overall sequence identity with the noncoding regions of RSV and AMPV. The role of these noncoding regions is likely the same as that for other paramyxoviruses, in which the gene end and gene start sequences control transcription termination and reinitiation, leading to a gradient of mRNA abundance that decreases from the 3⬘ end of the genome (N gene) toward the 5⬘ end (L gene) (136). The gene start consensus sequence of HMPV is CCCUGUUU/CA, and the start codon of each ORF is found 4 nucleotides downstream of this sequence (230). Using this knowledge, several noncoding regions have been duplicated in the HMPV genome to facilitate expression of green fluorescent protein and other heterologous ORFs, providing wonderful tools for HMPV research (24, 55). Steps in the HMPV replication cycle after the synthesis of RNA and viral proteins have not been investigated extensively. At present, as for other paramyxoviruses, it is assumed that

VOL. 24, 2011

HUMAN METAPNEUMOVIRUS

737

FIG. 3. Schematic representation of the HMPV life cycle. After attachment of the virion to the plasma membrane, the viral and plasma membranes fuse, resulting in uncoating of the virion and release of the RNP (containing the negative-sense viral RNA) into the cytoplasm. After primary transcription, the genome is replicated to produce the antigenome. The antigenome is used to synthesize genomic RNA, which is used to produce additional antigenomes for incorporation into progeny virions or as a template for secondary transcription. After translation, M proteins and RNPs are transported intracellularly to the plasma membrane and the viral glycoproteins F (fusion), G (glycoprotein), and SH (small hydrophobic) are transported from the endoplasmic reticulum (ER) to the Golgi apparatus and then the plasma membrane. Finally, new virions are assembled and are subsequently released from the plasma membrane by a budding process.

virus assembly and budding occur through similar mechanisms (136) (Fig. 3). EPIDEMIOLOGY Molecular Epidemiology and Virus Evolution When HMPV was first described as the causative agent of RTIs in children, it was immediately recognized that at least two genetic lineages of HMPV were circulating in humans

(231). Subsequent phylogenetic analysis of additional sequences obtained for the F and G genes revealed that each of these main lineages, A and B, can be divided into two sublineages: A1 and -2 and B1 and -2 (Fig. 4) (231). The maximum percent amino acid sequence identity between the F proteins of viruses belonging to lineages A and B was 95 to 97%; in contrast, there was only 30 to 35% identity for G protein sequences. The full-length genome sequences of prototypes of each of these lineages have become available in GenBank

738

SCHILDGEN ET AL.

CLIN. MICROBIOL. REV.

FIG. 4. Phylogenetic trees for fusion (F) and attachment (G) genes of selected HMPV isolates. For each of the four genetic lineages (233), four representative isolates were selected, and maximum likelihood trees were generated for the G gene (right) and for 451 nucleotides of the F gene (left). Numbers in trees represent percent amino acid identities between virus isolates.

(under accession numbers AF371337 [A1], FJ168779 [A2], AY525843 [B1], and FJ168778 [B2]) (54). The similarities between HMPV strains of different lineages are in the same range as that described for the subgroups of AMPV and RSV (204, 215). The circulation of the 4 genetic lineages of HMPV was confirmed in studies throughout the world, most notably in long-term retrospective studies conducted in the United States from 1981 to 2001 (248, 253). From these studies, it can be concluded that (i) the prevalence of particular lineages is not restricted to certain locations and times and (ii) multiple lineages can circulate in the same period at a given location. In 2004, a new variant of HMPV that was distantly related to previously described HMPV strains was detected in Germany (200). Unfortunately, the virus could not be isolated in cell culture and was therefore not characterized further, and other groups have not confirmed its detection. However, several other groups have subsequently reported the detection of newly emerging sublineages of lineages A and B (2, 9, 41, 44, 70, 118, 146, 242). In some of these studies, the available genetic information was limited to only small fragments of the HMPV genome, potentially giving rise to misclassifications. Nevertheless, it has become clear that sublineages of HMPV do not persist and that old lineages may be replaced with newly emerging variants. For instance, while lineage A1 circulated extensively in humans from 1982 to 2003 (233, 253), it has rarely been detected since 2004. In numerous other studies, it has been shown that the predominant circulating strains may vary by year and that predominant strains may be replaced, on average, every 1 to 3 years (2, 9, 40, 44, 70, 118, 140, 146, 152, 154, 166, 177, 185, 188, 242). Although antibody responses elicited against the highly con-

served F protein of HMPV may provide significant cross-protection against different HMPV lineages in animal studies (210), it has been postulated that antigenic variation may provide a plausible explanation for the cocirculation of multiple genetic sublineages of HMPV in humans (233). Virus neutralization assays performed with lineage-specific ferret antisera demonstrated that homologous virus neutralizing titers were significantly higher than titers against other HMPV lineages (233). Likewise, in reciprocal cross-neutralizing assays with sera from infected Syrian golden hamsters, the antigenic relatedness between viruses from two genetic lineages was relatively low (155). Based on these observations, as well as robust reinfections of cynomolgus macaques (232) and humans (66–68, 183, 253) with genetically distinct HMPV strains, it is possible that the cocirculation of multiple lineages of HMPV is facilitated by the limited cross-protection induced by HMPV. In this scenario, antigenic variation of the G and SH proteins, which may vary by as much as 70% at the amino acid level, may be sufficient to explain a selective advantage for heterologous virus lineages during subsequent epidemics, even in the presence of broadly cross-reactive anti-F humoral immunity in the population. With the advent of a rapidly increasing number of HMPV (and AMPV) gene sequences available in public databases, the evolutionary history and dynamics of HMPV have been studied. Investigations of the evolutionary dynamics of HMPV and AMPV-C, using G, F, and N nucleotide sequences, demonstrated higher substitution rates for the G gene (3.5 ⫻ 10⫺3 nucleotide substitution per site per year) than for the N (9 ⫻ 10⫺4 nucleotide substitution per site per year) and F (7.1 ⫻ 10⫺4 to 8.5 ⫻ 10⫺4 nucleotide substitution per site per year)

VOL. 24, 2011

HUMAN METAPNEUMOVIRUS

739

TABLE 2. Symptoms and clinical diagnoses associated with human metapneumovirus infection of children Valuea in reference: Parameter

No. of patients % of patients with symptom or diagnosis Fever Rhinorrhea Cough Wheezing Vomiting Diarrhea Rash Abnormal chest radiograph Bronchiolitis Pneumonia Croup Asthma Acute otitis media a

29

235

72

248

253

173

63

19

12

25

53

49

118

26

50

26

67 92 100 83 25 8 0 * 67 17 0 * 50

61 80 72 24 * * * 62 * * * * *

77 64 68 51 * * * 56 * * * * *

52 88 90 52 10 17 4 50 59 8 18 14 37

54 82 66 * 20 14 3 * * * * * 50

73 77 92 * * * * 85 23 23 * 27 15

44 90 90 56 36 14 2 67 48 34 4 * 6

13 2 16 6 4 1 5 * * 9 * * *

*, not reported.

genes (56, 259). Such high evolutionary rates are not uncommon for RNA viruses (120). In both studies (56, 259), a limited number of positively selected sites were found in the F gene, and none were found in the N gene. Mutations in the G gene were likely to be either neutral or positively selected. For the G protein of RSV, a strong association between neutralizing epitopes and positively selected sites has been reported (263). In contrast to the case for other paramyxoviruses, such as RSV, the HMPV G protein is not a major neutralizing or protective antigen (209). Presently, there is limited knowledge about the locations of neutralizing epitopes in the F protein of HMPV. It is possible that there is a correlation between positive selection of epitopes and neutralizing epitopes in the HMPV F protein, because the F protein represents the major neutralization and protective antigen. However, in contrast to those of other paramyxoviruses, the HMPV F gene does not display substantial evolutionary progressive drift (233). Extensive phylogenetic analyses have provided approximate calculations of the time of the most recent common ancestor of HMPV and AMPV-C. These analyses indicate that HMPV diverged from AMPV-C around 200 years ago (an average of 180 to 269 years, depending on the study and gene under investigation). The current genetic diversity of HMPV appears to have come about in the last ⬃100 years (97 years in reference 259 and 133 years in reference 56). Each of the main genetic lineages (A and B) appears to be 34 to 51 years old, while each of the sublineages (A1, A2, B1, and B2) appears to be less than 30 years of age (56, 259). Thus, the genetic diversity within the four sublineages is of extremely recent origin, with several lineage diversifications occurring at approximately the same time.

219, 223, 228, 238, 243). HMPV infections can occur throughout the year, but seasonality has been described in several studies, with the epidemiological peak occurring 1 to 2 months later than that observed for RSV epidemics (2, 3, 107, 157, 193, 245). The intriguing question of whether different HMPV lineages are associated with differences in clinical courses of disease has so far remained unresolved. Several groups have suggested that HMPV lineage A might be associated with more severe clinical disease (9, 126, 162, 240). However, others reported that lineage B may cause more severe illness (72, 185), while other groups found no evidence for differential severity caused by different HMPV lineages (4, 140, 160, 258). A better understanding of the host response to HMPV lineages is needed to understand the mechanisms that may contribute to differences in clinical severity. HMPV infections are observed in all age groups, with a high prevalence in pediatric patients (Table 1). The first HMPV infection appears to take place at 6 months of life, after which infections may occur repeatedly and frequently. The elderly represent the second group of patients that are severely affected by HMPV, and severe HMPV infections in the elderly occur despite high seroprevalence rates and independent of immunosenescence (62, 153). Reports on HMPV infections in otherwise healthy adults are relatively rare. Seroprevalence studies indicated that all adults have been infected with HMPV by the age of 25, with very high seroprevalence rates beginning from 5 years of age (68, 147, 150, 153, 255, 261, 262). The nosocomial impact of HMPV is estimated to be as high as that for RSV. In an HMPV outbreak in Japan, 34.8% of elderly patients who shared the same day care room in a hospital were infected with HMPV (116).

Clinical Epidemiology

CLINICAL FEATURES OF HMPV INFECTION

Since the first description of HMPV in 2001, the virus has been discovered worldwide on all continents and independent of the economic situations of different countries (1, 3, 5, 6, 17, 41, 44, 45, 49, 64, 67, 70, 83, 84, 93, 96, 97, 116, 132, 140, 145, 147, 148, 152, 154, 161, 165, 167, 170, 174, 181, 182, 186, 211,

Symptoms and Pathology HMPV is associated with a variety of symptoms and diagnoses localized to the respiratory tract (Table 2). Children with HMPV infection most commonly exhibit upper respi-

248/2,009 (1.4)

⬍3 and children of ⬎3

Nasopharyngeal swabs or aspirates NPA NPA NPA NPA Nasal washes Nasal washes NPA NPA NPA Nasopharyngeal secretions Nasopharyngeal swabs NPA NPA NPA Pharyngeal swab Throat swab NPA NPA Respiratory specimen (75% were nasal washes) Nasal washing

Respiratory samples

NPA NPA

Sample type

Hexamer PCR, RT-PCR, and cell culture

Singleplex RT-PCR RT-PCR of N gene RT-PCR of F gene RT-PCR of M and L genes RT-PCR of N and F genes Direct IF RT-PCR RT-PCR of M gene RT-PCR of N and F genes RT-PCR of F and N genes

GeneScan, RT-PCR of F gene RT-PCR of L gene Nested RT-PCR of matrix gene Multiplex RT-PCR, ELISA RT-PCR of F gene RT-PCR of L gene Multiplex PCR/RT-PCR RT-PCR of polymerase gene RT-PCR of N gene RT-PCR of M, F, and N genes

RT-PCR of nucleoprotein

RT-PCR of N gene RT-PCR of N gene, polymerase gene, and NL-N genes Pooling of clinical samples, nested RT-PCR

Detection method

1976–2001

October 2005–September 2007 April 2000–December 2007 October 2000–October 2007 October 2000–June 2005 January 2002–November 2003 December 2007–January 2008 December 2006–November 2008 December 2001–November 2004 October 2001–May 2004

January 2003–December 2006 2002–2006 October 2003–April 2004 2002–2006 October 2005–April 2007 November 2001–October 2002 2007–2009 June 2002–August 2002 2003 April–May 2002

October 2006–April 2007

July 2006–June 2008

May 2006–November 2007 October 2004–April 2008

Study period

ND

ND A1, A2, ND A1, A2, B1, B2 ND A2, A1, A1, A2, A1, A2,

B1 B1, B2 B1, B2

B1, B2

B1, B2

B1 (2004–2005), A2, B2 A (96%), B (4%) ND ND A2, B1, B2 ND B2, A2 ND ND ND

ND

A2, B1, B2

A2, B1, B2 ND

Genotypes detected

ND, not detected; NPA, nasopharyngeal aspirates; RT-PCR, reverse transcription-PCR; N, nucleocapsid protein/gene; F, fusion protein/gene; IF, immunofluorescence.

95/865 (11) 14/497 (2.8) 109/1,612 (6.8) 101/1,322 (7.6) 12/420 (2.9) 3/16 (18.8) 45/661 (6.8) 50/808 (6.2) 34/1,294 (2.6)

ⱕ3 ⱕ5 ⱕ2 ⱕ2 0–89 64–102 ⱕ14 Pediatric patients 0–ⱖ40

a

191/1,670 (11.4) 143/4,989 (2.8) 18/111 (16.2) 617/12,299 (5) 48/347 (13.8) 68/516 (13.2) 65/3,858 (1.7) 8/137 (5.8) 10/182 (5.5) 19/111 (17.1)

9/237 (3.8)

142/7,091 and 118/4,282

18/217 (8.3) 198/3,934 (5)

No. of HMPV-positive patients/total no. of patients (% positive)

ⱕ5 0–91 ⱕ1 0–16 ⱕ5 ⱕ5 ⱕ58 ⱕ14 ⱕ2 ⱕ3

Age groups are named A to K but are not specified in more detail 0–15

ⱕ2 Children

Age group (yr) in study

TABLE 1. Overview of selected recent clinical studies on the epidemiology and diagnostics of HMPV infectionsa

USA (248)

Alaska (208) Brazil (40) Austria (3) Spain (88) Peru (97) Australia (180) China (258) Italy (91) USA (96)

Brazil (177) Sweden (188) Spain (179) Germany (244) Italy (39) Israel (254) Cambodia (9) South Africa (79) Chile (151) Brazil (49)

Italy (73)

Scotland (90)

Uruguay (186) Switzerland (107)

Country (reference)

740 SCHILDGEN ET AL. CLIN. MICROBIOL. REV.

VOL. 24, 2011

ratory symptoms such as rhinorrhea, cough, or fever. Conjunctivitis, vomiting, diarrhea, and rash have been reported but are not frequent (234). The duration of symptoms prior to medical evaluation is usually less than a week, and limited data suggest that children shed virus for 1 to 2 weeks (67, 235, 248). Only one study has detected HMPV in serum by reverse transcription-PCR (RT-PCR) (159), suggesting that HMPV infection is usually limited to the respiratory tract. Studies of rodents and nonhuman primates have failed to detect HMPV in tissues outside the respiratory tract (7, 106, 131, 251, 257). The lower respiratory illnesses most frequently caused by HMPV are bronchiolitis, pneumonia, croup, and asthma exacerbation. Clinical signs and symptoms of HMPV infection overlap with those for other common respiratory viruses, and reliable distinctions cannot be made. Although symptoms usually overlap, differences in clinical presentation can occur. It has been reported that fever is more frequent in HMPV-infected patients, while rhinorrhea is observed more often in RSV-infected patients (19). Reinfection with HMPV occurs, although repeated infections are more likely to be limited to the upper respiratory tract in otherwise healthy children (248, 253). Some data suggest that dominant HMPV strains vary by season, presumably to avoid herd immunity, and thus reinfection may be more likely with heterologous viruses (2, 3). Further studies are needed to clarify the importance of antigenic variation of HMPV in human populations with respect to the clinical course of infection. HMPV is associated with acute otitis media, and viral RNA can be detected in middle ear fluid (199, 217, 252). Whether there is an association between HMPV infection and asthma is not clear. An Australian study of outpatient children with asthma did not identify an association between HMPV and asthma exacerbations (189), while a related study of outpatient children found a significant association between HMPV and the diagnosis of acute asthma (248). Studies of children hospitalized for wheezing and adults hospitalized with asthma exacerbations detected HMPV in a substantial number of these admissions (247, 250). Measurements of cytokines implicated in asthma pathogenesis in nasal washes of HMPV-infected infants have yielded conflicting results (119, 133). One retrospective study found a strong association between HMPV infection during infancy and the subsequent diagnosis of asthma (86). A major challenge for studies of respiratory viruses and asthma is the difficulty in firmly establishing a diagnosis of asthma during infancy, when acute wheezing associated with viral infections is common. However, the available data and analogy with RSV and human rhinoviruses suggest an association between HMPV and asthma exacerbations. One study reported histopathologic changes during HMPV infection of young patients. In this study, bronchoalveolar lavage (BAL) fluid samples and lung biopsy specimens from HMPV-positive children displayed epithelial cell degeneration or necrosis with detached ciliary tufts and round red cytoplasmic inclusions, hemosiderin-laden macrophages, frequent neutrophils, and mucus (237). However, these samples were obtained from patients with underlying disease, and similar studies of otherwise healthy children infected with HMPV have not been done. Studies of nonhuman pri-

HUMAN METAPNEUMOVIRUS

741

mates as well as small animals have demonstrated that HMPV infections remain restricted to the respiratory tract and do not spread to other internal organs. Histopathology studies of infected macaques and infected cotton rats have shown that infection is associated with a disruption of the epithelial architecture, sloughing of epithelial cells, loss of ciliation, and the presence of inflammatory infiltrates in the lungs (106, 131, 251, 257). HMPV-infected mice have been shown to develop parenchymal pneumonia and neutrophilic infiltrates during infection (50, 106). HMPV appears to exhibit a primary tropism limited to respiratory epithelia, as shown in immunohistochemistry studies of infected cynomolgus macaques, mice, and cotton rats. Viral expression was found in the epithelial cells of nasal tissue, all the way down to cells in the bronchioles, and was found less frequently in type I pneumocytes and alveolar macrophages (106, 131, 251).

Risk Groups Other than Children Populations at risk of HMPV infection are children, the immunocompromised, and the elderly. Most studies of the elderly were performed with study groups where the elderly were defined as persons above the age of 65 years (http://www .who.int/healthinfo/survey/ageingdefnolder/en/index.html). However, this definition is dependent on the geographic and social background of the population of the elderly. HMPV infection may be more severe in patients with underlying medical conditions. It has been shown that 30 to 85% of children hospitalized with HMPV have chronic conditions, such as asthma, chronic lung disease due to prematurity, congenital heart disease, or cancer (29, 63, 72, 168, 173, 225, 234). Hospitalization of adults or children for HMPV-associated lower respiratory tract infections is more likely for patients with underlying conditions such as asthma, chronic obstructive pulmonary disease (COPD), HIV infection, immunocompromised status, or prematurity (34, 66, 75, 78, 106, 122, 163, 168, 187, 193, 217, 229, 242, 253, 254). Immunosuppressed individuals. HMPV is capable of causing severe infections in immunocompromised hosts, a phenomenon that has been well described for most respiratory viruses, including influenza virus, RSV, and PIV. There are reports of fatal infection attributed to HMPV in cancer patients, and several studies suggest that HMPV is a relatively common cause of acute respiratory infection in children and adults with malignancy or hematopoietic stem cell transplants (27, 38, 80, 183, 249). The basis for the increased severity of disease in these different groups is likely related to a reduced capacity to control virus replication, but the mechanisms are not well understood. Long-term prospective studies are needed to better characterize disease due to HMPV infection in immunocompromised hosts. Older adults. Infections in adults are probably underreported, as many hospitals do not routinely screen adults for HMPV. The yearly incidence of HMPV infection in adults has been reported to be 4 to 11% (78, 247), but the incidence varies depending on the group studied, e.g., adults, high-risk patients, elderly adults, and residents of a long-term care facility. Infections in older adults are detected mostly in late winter and early spring, and viral coinfections are observed

742

SCHILDGEN ET AL.

mainly with RSV; up to 22.9% of infected elderly patients have been shown to be affected by at least one additional pathogen (78). HMPV is a significant cause of acute respiratory disease in older adults (⬎65 years) and adults with comorbid illness, such as COPD, asthma, cancer, or lung transplantation (30, 75, 103, 163, 229, 239, 241, 247). Given this, HMPV is responsible for many hospitalized cases (28, 30, 43, 241), and infections in the elderly can result in death (28, 30, 43). Pneumonia has been documented for 40% of HMPV-infected frail elderly subjects (28, 117, 149). It has been reported that the most frequent admission diagnoses for HMPV infection in the elderly are acute bronchitis, COPD exacerbation, pneumonia, and congestive heart failure (241). Twenty-seven percent of the hospitalized patients in this study had substantial airway infiltrates observed by chest radiographs, with an average length of hospitalization of 9 days, but the duration was twice as long for the high-risk group, reaching 34 days in the most severe cases, with 13.2% requiring intensive care. High fatality rates have been reported for elderly patients with underlying disease who died of general respiratory failure (33, 182, 247). In one study, a case of HMPV-induced fatal pneumonia in an old woman who had no medical condition other than advanced age was presented (30), illustrating that HMPV infection can cause severe pneumonia leading to death in otherwise healthy elderly individuals. In addition, a single case of severe acute pericarditis associated with HMPV was reported for an otherwise healthy 62-year-old woman (52); thus, complications may occur that might be linked to HMPV infection. The reasons for the higher morbidity in older adults have not been determined and require attention. Increased morbidity as well as a delay in clearance of symptoms of virus infections has been reported for the elderly, a feature consistent with the impairment of innate and adaptive immunity commonly associated with aging. Aging causes both qualitative and quantitative alterations, and intrinsic defects in the T cells of aged mice have been shown to contribute to decreased virus-specific T cell responses (33, 121–123, 144, 187). Although defects in the early events after virus infection may also influence the ageassociated delay in clearance of the virus, it is possible that one reason for the clinical aggravation observed in the elderly could be an exaggerated immune response with inflammation, as opposed to a declining immune response (214). In the context of virus infection, these outcomes may be affected differently by the strain or HMPV subtype. It is clear that additional studies are required to determine the nature of the age-related defects during HMPV infection, and appropriate animal models are essential for these investigations. Healthy adults suffer from asymptomatic infection, colds, and influenza-like illnesses (102). Asymptomatic infections are more common for HMPV infections than for RSV or influenza A virus infections. Asymptomatic infection in adults was also reported in a survey study of immunocompromised bone marrow transplant recipients (53). Both asthma (75, 247) and COPD (6, 9, 17, 20, 74, 103, 115, 118a, 163, 179, 194, 214) are exacerbated in immunocompromised adults. Serious and prolonged respiratory infections are associated with mortality in adults with underlying disease or hematological malignancies (247) and following hematopoietic stem cell or lung transplantation (139, 191, 216). Elderly subjects frequently suffer from

CLIN. MICROBIOL. REV.

bronchitis and pneumonia, and HMPV is responsible for many hospitalized cases in this population (17, 115, 128). Coinfections Many studies evaluating HMPV infection have tested for other viruses by using sensitive methods and have detected RSV in 5 to 17% of patients infected with HMPV (10, 17, 29, 31, 38, 49, 63, 67, 72, 81, 84, 119, 133, 159, 168, 173, 182, 189, 225, 235, 238, 248, 253). Most studies have not described exacerbated disease in patients with codetection of multiple viruses. Note that highly sensitive RT-PCR techniques may detect viruses for several weeks after an acute infection. A few studies of hospitalized patients have described much higher coinfection rates (30 to 60%) (87, 119, 130, 238), raising the question of whether HMPV infections are more severe if another virus is present. One group addressed this question by using a nested RT-PCR assay to test BAL fluids from 30 intubated infants with RSV infection, and they detected HMPV in 21/30 (70%) infants (98). They subsequently used the same nested PCR assay to test specimens from children admitted to the intensive care unit and the general wards. HMPV and RSV coinfections were detected in 18/25 (72%) intensive care patients and in 15/171 (9%) general ward patients, leading the authors to conclude that dual infection with RSV and HMPV was associated with severe bronchiolitis (205). However, a study of 46 inpatients with either mild or severe RSV disease found no coinfections with HMPV (141). Also, a Dutch study did not find any HMPV-RSV coinfections in children with severe RSV bronchiolitis (236). Whether these conflicting findings are due to methodological differences or to variability in circulating viruses is unknown. Further studies are needed to clarify the nature of disease associated with codetections; however, it is clear that the majority of HMPV infections are not associated with other viruses and that HMPV is a primary respiratory pathogen. Treatment Options The majority of children infected with HMPV can be managed with supportive care. For infants and children who require hospitalization, the primary therapies are supplementary oxygen and intravenous hydration. Bronchodilators and corticosteroids have been used empirically, but there are no controlled trials of these medications for HMPV and no data to support or refute their efficacy. One animal study suggested that there were benefits of bronchodilator and corticosteroid treatment of experimental HMPV infection in cotton rats (104). The only currently licensed antiviral drug for a related virus, i.e., RSV, is ribavirin, a nucleoside analogue administered by aerosol. Both ribavirin and polyclonal human immunoglobulin exhibit in vitro neutralizing activity against HMPV equivalent to their activity against RSV (256). There are no published animal or human data for these interventions, although they may be worthy of consideration for severely immunocompromised hosts with lower respiratory infections due to HMPV. Ribavirin has been used in severely immunocompromised patients, such as hematopoietic stem cell transplant recipients, often in conjunction with RSV-specific immunoglobulin, with some evidence of efficacy (60).

VOL. 24, 2011

HUMAN METAPNEUMOVIRUS

IMMUNE RESPONSES IN HUMANS AND ANIMALS Innate Immunity Acute viral infection is known to induce an early innate immune response characterized by induction of an antiviral type I interferon (IFN) response that involves JAK/STAT signaling, leading to the production of a variety of antiviral proteins, particularly IFN-stimulated gene products (218). Tolllike receptors (TLRs) and RNA helicases (RIG-I and MDA-5) are pattern recognition receptors most commonly activated by viral infections, and their activation contributes to a signaling cascade that governs the expression of proinflammatory and immune mediators. The TLRs, some of which are located in the endosomal cellular compartment, operate mainly in primary antigen-presenting and dendritic cells, while RIG-I and MDA-5 are cytoplasmic sensors and have been identified as being essential for IFN induction by several viruses. Mitochondrial antiviral signaling protein (MAVS) is an adaptor protein linking both RIG-I and MDA-5 to downstream activation of IRF3 and NF-␬B, ultimately leading to type I IFN and IFNstimulated gene expression (127, 206). RNA viruses use a large number and variety of mechanisms to subvert innate antiviral host defenses, and many of these mechanisms involve evasion of IFN and IFN-regulated responses (89). Limited information is available on the interaction of HMPV and the innate immune system. In cultured human epithelial cells, HMPV was shown to be a strong inducer of the type I IFN pathway (14), and HMPV sensing and subsequent activation of the IFN pathway appeared to occur through the RIG-MAVS pathway, with no involvement of MDA-5 (143). Different HMPV proteins have been suggested to subvert the antiviral IFN pathway. Mutant HMPV lacking the G gene was shown to induce higher levels of chemokines and type I IFN in infected airway epithelial cells than those induced by the wildtype virus (12, 13, 15). The phosphoprotein in HMPV genotype B viruses was shown to be an IFN antagonist, but interestingly, the phosphoprotein in genotype A HMPV lacked this activity (95). In this study, it appeared that RIG-I signaling was not blocked by the phosphoprotein, while other studies suggested that the G protein binds to RIG-I to inhibit the IFN pathway (12, 13, 15). Two studies have shown that HMPV blocks double-stranded RNA (dsRNA)-induced IFN expression (100, 101), and in cultured epithelial cells, HMPV prevented IFNinduced phosphorylation and subsequent nuclear translocation of STAT1 (61). These findings contradict a report showing that HMPV infection of human A549 cells induces STAT1 phosphorylation (14). The contradictory results might relate to differences in the strains of HMPV used in the studies, as accumulating evidence suggests that there are strain differences in the host response to HMPV infection. Limited data are available on the innate immune response to HMPV infection in humans; however, the available data suggest that differences may exist between HMPV and RSV. A Finnish study reported increased levels of interleukin-8 (IL-8) and decreased levels of CCL5 (RANTES) in nasal secretions of HMPV-infected children compared with levels from RSVinfected children (119). In contrast, a prospective study of infants infected with HMPV revealed significantly lower levels of IL-1␤, IL-6, IL-8, IL-12, and tumor necrosis factor alpha

743

(TNF-␣) in nasal washes than those in RSV- or influenza virus-infected infants (133). Experimental HMPV infection of human dendritic cells ex vivo also induced cytokine profiles that differed significantly from responses evoked by RSV (100). In addition, a study with HMPV-infected BALB/c mice demonstrated significantly larger amounts of IFN-␣ than those in mice infected with RSV (99). These findings suggest that HMPV may induce a distinct host response, perhaps characterized by potent innate responses to infection. Humoral Response The prevalence of HMPV antibodies in persons between the ages of 6 months and 1 year, as measured by immunofluorescence assay, is 25%, and this prevalence increases to 55% for those between 1 and 2 years, to 70% for 2- to 5-year-olds, and to 100% for those over 5 years of age. Neutralizing antibody titers are lower for younger than for older children, with proportions of 25% for 6- to 12-month-olds, 31% for 1- to 2-yearolds, 38% for 2- to 5-year-olds, and 75 to 100% for those over 5 years of age (231). Thus, while all persons have been infected by age 5, a substantial proportion of those who are seropositive by immunofluorescence assay do not exhibit high neutralizing antibody titers. Consistent with these findings, several studies have shown that high HMPV seropositivity is associated with age and that high overall rates of seropositivity occur after the age of 5 years (68, 142, 181, 255). In general, it appears that despite protective antibody titers in adults, reinfections with homotypic and heterotypic HMPV strains occur in both healthy and immunocompromised humans (66, 157, 183, 248, 253). It is not known whether this phenomenon is due to limited cross-protective immunity between strains of HMPV, but these findings have led to the hypothesis that humoral immune responses may have only a minor role in the clearance of HMPV. However, this hypothesis is weakened by the observation that all permissive animal models, i.e., those that can be infected with HMPV and suffer from HMPV-associated disease, have been shown to develop serum neutralizing antibodies upon experimental HMPV infection (7, 106, 155, 232, 251, 257). Furthermore, passive antibody transfer alone, either polyclonal or monoclonal, has been shown to protect against HMPV replication and disease (7, 227, 246). Several different experimental approaches have shown that the HMPV F protein is a primary target of neutralizing antibodies (25, 171, 196, 209). However, since virtually all humans are seropositive yet severe infection occurs, antibody-mediated protection is probably not sufficient to prevent disease pathogenesis. One study using macaques showed waning immunity at 12 weeks and complete loss of protection 8 months following experimental infection (232). A poor HMPV antibody response was observed in aged mice, and this was not due to less viral replication, as the virus load in the lung at day 3 was shown to be greater in the older mice than in younger mice (51). This impaired antibody response was reflected in the IgG2a isotype, where the production of neutralizing antibodies was also impaired. It is possible that the HMPV-specific antibody response may contribute to protection against the severity of illness. This hypothesis is consistent with the finding that passive transfer of HMPV-immune serum protects naïve BALB/c mice from challenge and with results showing that a neutral-

744

SCHILDGEN ET AL.

izing monoclonal antibody to the F protein confers protection against challenge (8, 246).

Cellular Immunity Reports of severe and fatal infections in immunocompromised populations suggest that T cell immunity is important for virus clearance and resolution of illness (38, 69, 139, 164, 183, 249). Many of these reports identified lymphopenia and cytotoxic therapy as important risk factors for severe HMPV disease, emphasizing a likely role for T cells in protection. Consistent with these findings, HMPV infection has been shown to be more severe in immunodeficient HIV-infected persons (85, 118a, 156, 157). HLA class I-restricted cytotoxic T lymphocyte (CTL) responses have been evaluated ex vivo in cells from HMPV-infected humans by enzyme-linked immunosorbent spot (ELISPOT) assay and chromium release assays (109). However, the contributions of T cells during primary and secondary infections of humans are unclear. In mice, depletion of NK cells was associated with significantly increased HMPV titers in the lungs, suggesting that cytotoxic cell types may contribute to HMPV immune surveillance and control (7). The contribution of T cells to HMPV immunity has been investigated to a limited extent by use of animal models. CD4⫹, CD8⫹, or CD4⫹ CD8⫹ T cell-depleted mice exhibited diminished weight loss and lung histopathology, suggesting that host immunity, while required for virus clearance, contributes to disease (129). However, if secondary virus challenge was performed in T cell-depleted mice, no difference in titer occurred between depleted and control groups of mice at the single time point examined, despite an absence of antibody in the CD4⫹ T cell-depleted mice. Therefore, the contributions of CD4⫹ and CD8⫹ T cells to protection or pathology remain poorly defined. Murine major histocompatibility complex (MHC) class I-restricted epitopes for HMPV have been described (108, 169). One group used algorithms to identify candidate epitopes and tested predicted epitopes by peptide immunization (108). This group found that a few of these epitopes induced peptidespecific CTL responses against the same immunizing peptide and resulted in modest protection against virus challenge. However, the peptides were not screened against CTLs induced by virus infection, and thus it is not clear that these are true epitopes presented during natural infection. A similar bioinformatic approach was used to identify potential H-2drestricted epitopes, i.e., epitopes recognized by murine MHC II, the analog of the human leukocyte antigen (HLA) complex (169). Predicted epitopes were screened as synthetic peptides against HMPV-immune splenocytes from BALB/c mice by ELISPOT assay. These experiments identified H-2d-restricted dominant (M2181–189) and subdominant (N307–315) epitopes. Primary T cell lines generated ex vivo by stimulation with these peptides were CD8⫹, IFN-␥ secreting, and functional for lytic activity. Adoptive transfer of these cell lines into Rag⫺/⫺ mice provided a modest reduction in virus titer, but lung pathology or signs of disease were not determined. These studies suggest that T cells contribute to protective immunity.

CLIN. MICROBIOL. REV.

ANIMAL MODELS OF HMPV INFECTION Several animal models have been developed to study HMPV infections. These animal models include mice, cotton rats, hamsters, guinea pigs, and ferrets, as well as nonhuman primates such as chimpanzees, macaques, and African green monkeys (155, 201). Each animal model has unique features that allow for investigations into the mechanisms of immunity and disease pathogenesis. The following sections summarize the roles of animal models and their contributions to our understanding of the biology of HMPV infection, replication, and disease intervention strategies. Mouse Model Experimental HMPV infection of mice (Mus musculus) has been studied with several mouse strains, but the majority of these studies were conducted with the BALB/c strain, as numerous studies have demonstrated robust HMPV replication in the lungs of BALB/c mice, with peak titers from days 5 to 7 (7, 8, 51, 213). In an early study, HMPV was shown to exhibit biphasic growth kinetics in the lungs of BALB/c mice, with peak titers occurring at days 7 and 14 postinfection (p.i.), while infectious HMPV could be recovered from the lungs until day 60 p.i. and HMPV RNA could be detected at ⱖ180 days p.i. (7). These results suggested the ability of HMPV to establish persistence following infection and are consistent with a more recent publication showing persistence of HMPV RNA in the lungs of mice, with associated pulmonary inflammation, out to day 154 p.i. (105). However, similar results were not reported in other studies showing that HMPV induces a self-limiting infection. The observed biphasic growth kinetics might reflect the specifics of the virus strain used or methods employed. The level of illness associated with weight loss has been linked with the level of virus replication and ranges from 5 to 20% weight loss, depending on initial viral inoculums (7, 8, 213). No viral RNA or infectious virus has been detected in the serum, spleen, kidneys, heart, trachea, or brain tissue (7). However, it has also been reported that HMPV has limited replication in mice (155). In these studies, BALB/c mice infected intranasally with HMPV/NL/1/00 were shown to be semipermissive for HMPV infection. The disease pathogenesis associated with HMPV infection has been well characterized in mice. In general, following intranasal HMPV infection, lung histopathology reveals a prevalent mononuclear cell infiltration in the interstitium, beginning at day 2 p.i. and decreasing by day 14 p.i., and this has been associated with airway remodeling, increased mucus production, and airway hyperresponsiveness (AHR) (7, 129). In the BALB/c mouse model of HMPV infection, infection has been associated with an indolent inflammatory response characterized by innate immune and CD4⫹ T cell trafficking to the lungs, low-level IFN expression, induction of IL-10 expression at later stages of infection, and delayed CTL activity that coincides with persistent virus replication in the lungs (8). In addition, one study showed age-associated aggravation of HMPV clinical disease in BALB/c mice (51). Young and aged mice showed respiratory dysfunction, weight loss, and similar histopathological abnormalities. However, aged mice were far more susceptible than young mice to HMPV infection. It was

VOL. 24, 2011

HUMAN METAPNEUMOVIRUS

suggested that this increased susceptibility was linked to the loss of cellular immune responses that are important for controlling HMPV infection. The virus and host mechanisms that contribute to pathogenesis are not fully understood, but several studies have addressed these. In an intranasal inoculation model, it was shown that HMPV-infected BALB/c mice developed parameters of clinical disease, including airway obstruction and hyperresponsiveness, and that this was mediated predominantly by CD4 T cells in comparison to CD8 T cells (129).

Cotton Rat Model The cotton rat (Sigmodon hispidus and Sigmodon fulviventer) has become an accepted model for studying respiratory virus infections. Numerous studies have used the cotton rat model to evaluate HMPV infection, vaccine candidate efficacy, and disease pathogenesis (25, 35, 111, 155, 184, 197, 209, 210, 222, 251). In cotton rats infected intranasally with HMPV, peak levels of virus replication in the lung generally occur between days 4 and 5 p.i. Clinical symptoms are generally not observed, although infection has been associated with significant inflammation and histopathological changes associated with the development of peribronchial inflammatory infiltrates in the lower respiratory tract (251). No significant peripheral tissue pathology was evident in any of multiple tissues examined, and immunostaining revealed the presence of antigen at the apical surface of respiratory epithelial cells, suggesting a tropism for respiratory epithelial cells. Differences in susceptibility to different HMPV strains have been examined. In one study, cotton rats were inoculated intranasally with one of three strains, i.e., HMPV-26575, HMPV-26583, or RL Bx (257). Genetic analysis indicates that HMPV-26575 belongs to genotype B, while HMPV-26583 and RL Bx belong to genotype A (257). Interestingly, the genotype A viruses had similar replication kinetics and grew to high titers; however, the type B virus failed to replicate to high levels in the lungs of cotton rats. Increasing evidence demonstrates that the results obtained were more likely to be due to strain differences than genotype differences. Young adult cotton rats are good models for evaluating disease intervention strategies for HMPV, because changes seen in cotton rats appear to be similar to those reported to occur in nonhuman primates following HMPV infection (131). In both species, HMPV infection results in inflammation in and around the bronchi and bronchioles, with markedly increased numbers of leukocytes surrounding these regions. Virus-specific antibody staining revealed HMPV antigen predominantly on the apical surface of the columnar cells, and peak viral titers in the lungs generally occurred around day 5 p.i. (201). Infection is accompanied by an inflammatory response characterized by upregulation of several chemokines and cytokines in the lungs, including IFN-␣, CCL5 (RANTES), CCL2 (MCP-1), CCL3 (MIP-1␣), and IL-2 mRNAs (32). HMPV infection in cotton rats results in a partially protective immune response, a reduced level of virus replication in the lungs following challenge, and a virus-specific serum neutralizing antibody response (32).

745

Hamster and Ferret Models Several studies have evaluated HMPV infection and vaccine efficacy in hamsters (Mesocricetus auratus). HMPV replication in the lungs of hamsters has been shown to be high (105 PFU/g lung tissue) and similar to levels observed in cotton rats (155). This feature makes the hamster an ideal model for evaluating vaccine antigenicity and efficacy (113). There has been little done in the ferret model of HMPV infection. In one study that evaluated a panel of small animal models for HMPV, ferrets were infected intranasally with 106 PFU of HMPV, and at day 4 p.i., levels of virus in the turbinates and lung tissues were determined (155). The results showed that like hamsters, ferrets support HMPV replication in the respiratory tract to high titers, e.g., 4 to 4.7 log10 PFU/g tissue. Interestingly, there was no evidence of fever in the ferrets, which were monitored daily, and no clinical signs of illness. However, both hamsters and ferrets developed neutralizing HMPV antibodies. Given these limited features, the higher costs associated with ferrets, and the limited availability of immunological reagents for ferrets, it is likely that few studies will emerge using the ferret model. HMPV Infection in Nonhuman Primates To date, several nonhuman primate species have been evaluated for HMPV replication and viral pathogenesis, including rhesus macaques, cynomolgus macaques, African green monkeys, and chimpanzees (22, 155, 210, 221). In chimpanzees screened for the presence of neutralizing anti-HMPV antibodies in serum, 61% (19 of 31 animals) of animals were seropositive for either genotype A or genotype B HMPV strains (210), i.e., HMPV circulated within the chimps’ group or was transmitted by their keepers. When chimpanzees were infected experimentally with HMPV, they demonstrated signs of clinical disease, including nasal discharge, thick mucus, and decreased appetites; however, despite these clinical changes, viral titers reached only 1.8 to 3.2 log10 in either the lower or upper respiratory tract (210). Rhesus macaques infected with 105.2 50% tissue culture infective doses (TCID50) of HMPV demonstrated only low levels of viral replication, with peak titers ranging from 0.9 to 2.6 log10 TCID50 in nasopharyngeal swabs and tracheal lavage fluid. In contrast, rhesus macaques infected with a similar inoculum yielded viral titers of 2.2 to 3.7 and 4.9 to 5.0 log10 TCID50 from the upper and lower respiratory tracts, respectively, and developed serum neutralizing antibody titers ranging from 9.1 to 10.9 log2, with initial infection conferring resistance against subsequent infections with both homologous and heterologous HMPV strains (155, 221). The African green monkey model has been utilized to investigate the generation of novel HMPV-based vaccines. VACCINE DEVELOPMENT A number of vaccines against HMPV have been investigated, including those prepared from chimeric viruses, liveattenuated viruses, and subunits of the virus. One of the first vaccine studies evaluated the efficacy of a chimeric virus vector consisting of bovine parainfluenza virus type 3 (PIV3) containing the F and HN genes of human PIV3 and expressing the HMPV F protein (222). In this study, vaccination of hamsters

746

SCHILDGEN ET AL.

or African green monkeys induced HMPV-specific neutralizing antibodies that protected against HMPV challenge (221, 222). Other chimeric vaccine candidates were evaluated, such as a virus where the nucleoprotein or phosphoprotein of HMPV was replaced with that of AMPV type C. High levels of protective neutralizing antibodies were observed following intranasal vaccination, and upon challenge, lung virus titers were reduced substantially compared to those of controls at 3 days postchallenge (184). A study examining a chimeric virus vaccine based on alphavirus replicon particles encoding the G or F protein of HMPV demonstrated that the virus harboring the F protein of HMPV induced protective immunity against subsequent challenge. The latter observation made for HMPV is in contrast to the case for alphavirus harboring the G protein, which did not appear to be immunogenic and protective (170). A live-attenuated HMPV vaccine candidate was generated by repeated passages of HMPV at low temperature in Vero cells, resulting in the accumulation of mutations in the viral genome (111). These mutations were reverse engineered into a wild-type HMPV backbone and resulted in a number of viruses with a temperature-sensitive phenotype. Replication of these temperature-sensitive HMPV vaccine candidates was reduced in the upper respiratory tract of hamsters and undetectable in the lower respiratory tract, but it was sufficient to induce high titers of protective HMPV-specific antibodies (111). Other live-attenuated vaccine candidates were generated by deleting the SH and/or G gene or the second ORF of the M2 gene. These vaccine candidates were shown to be attenuated in hamsters and nonhuman primates but were immunogenic, as they protected animals from subsequent challenge (25, 35, 197). However, a mutant virus lacking the M2-1 gene did not induce protective immunity, indicating that M2-1 is essential for virus replication. The F protein has been used to develop several subunit vaccine candidates. Vaccination of hamsters with adjuvanted soluble F protein was shown to induce protective immunity against HMPVs of both genetic lineages (112). In addition, DNA vaccination with plasmid DNA carrying the F gene or vaccination with a purified soluble F protein lacking the transmembrane domain induced protective immunity (48). Moreover, the use of CTL peptide epitopes has been tested as a potential vaccination strategy in BALB/c mice: vaccination with such peptides was shown to reduce viral loads and immune pathology in the lungs of HMPV-challenged mice, an effect linked to enhanced expression of Th1 cytokines, including IFN-␥ and IL-12 (108). Venezuelan equine encephalitis virus replicon particles encoding the HMPV F or G protein have also been evaluated for immunogenicity and protective efficacy in mice (171). Although several vaccine strategies have been developed, caution is needed, based on the disastrous outcome of formalin-inactivated RSV (FI-RSV) vaccines. FIRSV vaccines administered to young children in the late 1960s sensitized vaccinees for vaccine-enhanced illness upon natural RSV infection (26). Similar FI-HMPV vaccines have been examined in mice, and the results show that HMPV challenge of FI-HMPV-vaccinated mice also leads to vaccine-enhanced disease (59, 260).

CLIN. MICROBIOL. REV.

DIAGNOSTICS Molecular Diagnostics Currently, there exists neither a “gold standard” nor a consensus assay for the detection of HMPV in clinical samples. RT-PCR-based techniques are generally the methods of choice for the detection of HMPV (29, 71, 76, 98, 158, 182, 230); however, other assays, such as isothermal real-time nucleic acid sequence-based amplification (NASBA), have been used (92). Most PCR protocols detect all HMPV genotypes and rely on conserved and essential regions within the N gene or the F gene. Commercial singleplex assays are available from several diagnostic companies. These assays include both RT-PCR (e.g., assays by GenProbe/Prodesse, San Diego, CA) (82) and NASBA (bioMe´rieux, Marcy l’Etoile, France) (92) assays. Unfortunately, data on the utility of those assays are rare, despite the fact that NASBA was shown to be as sensitive as PCRbased detection methods (92). Recent studies used multiplex assays to evaluate large cohorts of patients coinfected with two or more pathogens (21, 190). In some patients, up to five pathogens were detected simultaneously. Most multiplex assays for the detection of respiratory viruses also include HMPV detection reagents. The xTAG respiratory virus panel (RVP; Luminex, Toronto, Canada) was shown to have superior negative predictive values, with acceptable positive predictive values that were marginally lower than the positive predictive value of the ResPlex II test by Qiagen (Hilden, Germany) (11). The ResPlex II assay has a broader pathogen range and detects up to 19 different viruses, compared to 17 viruses detected by the xTAG RVP assay (11). With respect to HMPV, the xTAG RVP, ResPlex II, and MultiCode-PLx assays have similar detection efficiencies (11). In addition, the MultiCode-PLx assay (EraGen Biosciences, Madison, WI) (175) has an identical negative predictive value to that of the Qiagen assay, but both values are lower than that for the Luminex assay (11). Thus, the xTag assay has received FDA approval, which is a prerequisite for its favored use in routine laboratory diagnostics in the United States, as in-house validation would otherwise be required. Another FDA-approved multiplex assay is the FilmArray respiratory panel by Idaho Technology Inc. (Salt Lake City, UT). This assay is a fully automated cassette that contains all necessary reagents, starting from extraction via a combined multiplex RT-PCR followed by singleplex second-stage PCRs to the final detection via endpoint melting curve. The assay is designed to detect up to 15 different agents in a single sample and has a very high specificity (99.2% for HMPV). An alternative multiplex detection method for respiratory pathogens is the RespiFinder technology (Pathofinder, Maastricht, The Netherlands), which is based on multiplex PCRs that are analyzed by subsequent capillary gel electrophoresis (192). This technology make use of multiplex ligation-dependent probe amplification (MLPA) and employs two probes which ligate exclusively in the presence of target-specific complementary sequences (34, 192). This assay was shown to have high sensitivity (98.2%) and specificity (100%) for HMPV in a study that investigated 144 clinical samples (192). Unfortunately, this assay is restricted in its use to those laboratories that are equipped with a capillary sequencing unit. The first version of this assay detected up to

VOL. 24, 2011

HUMAN METAPNEUMOVIRUS

15 pathogens simultaneously, but the second version, i.e., RespFinder19, is able to detect 19 pathogens simultaneously, including 15 respiratory viruses. Recent studies have shown that the RespiFinder assay (34) and the Seeplex RV15 ACE assay (Seegene, Eschborn, Germany) (65) are more sensitive than cell culture but comparable to singleplex real-time RT-PCR. There are currently other novel technologies being developed for respiratory virus diagnostics. Some examples include microfluid chip-compatible assays (36), RTPCR coupled to electrospray ionization mass spectrometry (42), and surface-enhanced Raman spectroscopy assays (207), which may improve the sensitivity and range of pathogen detection.

Fluorescent-Antibody Staining and ELISA Diagnosis of HMPV infection is based on the direct detection of viral components (protein, particles, or RNA) rather than indirect detection of antiviral antibodies in patient sera; however, there is value in evaluating antibody levels, particularly for understanding vaccine efficacy. Several studies have reported high seroprevalences based on enzyme-linked immunosorbent assays (ELISAs) (68, 145, 176, 262) or microneutralization assays (77, 153). ELISA detection of HMPV is generally performed with in-house assays, because assay kits are currently not commercially available. As an alternative method, cytospin-assisted direct immunofluorescence assay (DFA) is used occasionally for HMPV diagnosis in Europe and has become quite standard in the United States, as it allows for rapid detection of viral proteins from clinical samples, with acceptable sensitivities (124, 137). Commonly used antibodies are those from Chemicon/Millipore (Chemicon International, Temecula, CA) (137, 138); these monoclonal antibodies are offered as complete kits, e.g., the Light Diagnostics human metapneumovirus (HMPV) direct immunofluoresence assay, or as the SimulFluor HMPV/RSV reagent (both from Millipore, Billerica, MA). Also, Diagnostic Hybrids (Athens, OH) has developed and launched FDA-cleared assays, namely, the D3 DFA metapneumovirus identification kit for the detection of HMPV and the D3FastPoint L-DFA respiratory virus identification kit, which allows the identification of 8 different viruses, including HMPV. The latter test is claimed to be as sensitive and accurate as DFA, but a recent study showed that it is less sensitive than PCR and DFA; however, the time for multiplex detection is shorter (M. Barger, D. Vestal, M. Nye, and B. A. Body, presented at the 26th Clinical Virology Symposium, 25 to 28 April 2010, Daytona Beach, FL).

Cell Culture HMPV can be cultured in several cell lines, including tertiary monkey kidney cells (231), Vero cells (231), LLC-MK2cells, BEAS-2B cells (226), A549 cells (14), and HepG2 cells (202). These cell culture models facilitate HMPV research; however, there are caveats related to the different envelopes the viruses acquire as they bud from the cells, and these different envelopes may modify immune responses and may interfere with some assays.

747

FUTURE PERSPECTIVES Although a substantial amount of knowledge on HMPV has been gained during the last decade, many issues remain unsolved. Despite extensive efforts to understand the molecular basis of the HMPV life cycle, the functions of several HMPV proteins need to be investigated further. For instance, studies regarding the F protein have shown that, as for most paramyxoviruses, the trigger that leads to the membrane fusion event remains unknown. The binding of F protein to integrin ␣v␤1 may provide such a trigger and is an intriguing area for further investigation. In addition, the function of the SH protein of paramyxoviruses has remained elusive, and more work is needed to gain detailed knowledge of the HMPV polymerase complexes and the roles of M2-1 and M2-2 during virus replication. As for the clinical impact of HMPV, long-term studies are needed to characterize disease pathogenesis and to understand immunity in very young, old, or immunocompromised hosts responding to HMPV infection. There is also a need to optimize commercial diagnostic reagents and methods for detection of HMPV infection. Although a plethora of studies have been done on the clinical impact of HMPV, more detailed studies are needed to clarify the importance of antigenic variation of HMPV in human populations. This is important for the development of a cross-protective vaccine and for understanding antiviral approaches. Moreover, there is a need to develop safe and effective vaccines that induce protective immunity and perhaps give cross-protection against related viruses, such as RSV. To achieve these goals, a better understanding of the host response is needed at the genome level as well as the immune level, and the mechanisms of innate immunity that facilitate adaptive immunity need to be determined. ACKNOWLEDGMENTS V.S., O.S., C.M., and R.A.T. have no conflicts of interest to declare. B.V.D.H. is named inventor on several patents related to HMPV. These patents have been licensed to MedImmune USA. R.F. is named inventor on several patents related to HMPV. These patents have been licensed to MedImmune USA. R.F. is also a holder of certificates for 1% of the shares in ViroClinics Biosciences B.V. ViroClinics Biosciences is a contract research organization providing an array of services to the biopharmaceutical industry. To avoid any possible conflict of interests, Erasmus MC policy dictates that the shares as such are held by the Stichting Administratiekantoor Erasmus Personeelsparticipaties. The board of this foundation is appointed by the Board of Governors of the Erasmus MC and exercises all voting rights with regard to these shares. J.W. served as a consultant to MedImmune and Novartis and also serves on the Scientific Advisory Board of Quidel. R.A. is an employee of the U.S. Government. This work was prepared as part of official duties. The views expressed in this article are those of the authors and do not necessarily reflect the official policy or position of the Department of the Navy, the Department of Defense, or the U.S. Government. REFERENCES 1. Abdullah Brooks, W., et al. 2007. Human metapneumovirus infection among children, Bangladesh. Emerg. Infect. Dis. 13:1611–1613. 2. Aberle, J. H., S. W. Aberle, M. Redlberger-Fritz, M. J. Sandhofer, and T. Popow-Kraupp. 2010. Human metapneumovirus subgroup changes and seasonality during epidemics. Pediatr. Infect. Dis. J. 29:1016–1018. 3. Aberle, S. W., J. H. Aberle, M. J. Sandhofer, E. Pracher, and T. PopowKraupp. 2008. Biennial spring activity of human metapneumovirus in Austria. Pediatr. Infect. Dis. J. 27:1065–1068. 4. Agapov, E., K. C. Sumino, M. Gaudreault-Keener, G. A. Storch, and M. J. Holtzman. 2006. Genetic variability of human metapneumovirus infection: evidence of a shift in viral genotype without a change in illness. J. Infect. Dis. 193:396–403.

748

SCHILDGEN ET AL.

5. Albuquerque, M. C., et al. 2009. Novel respiratory virus infections in children, Brazil. Emerg. Infect. Dis. 15:806–808. 6. Ali, S. A., et al. Human metapneumovirus in hospitalized children in Amman, Jordan. J. Med. Virol. 82:1012–1016. 7. Alvarez, R., K. S. Harrod, W. J. Shieh, S. Zaki, and R. A. Tripp. 2004. Human metapneumovirus persists in BALB/c mice despite the presence of neutralizing antibodies. J. Virol. 78:14003–14011. 8. Alvarez, R., and R. A. Tripp. 2005. The immune response to human metapneumovirus is associated with aberrant immunity and impaired virus clearance in BALB/c mice. J. Virol. 79:5971–5978. 9. Arnott, A., et al. 1 February 2011. Genetic variability of human metapneumovirus amongst an all ages population in Cambodia between 2007 and 2009. Infect. Genet. Evol. 10. Bach, N., et al. 2004. Acute respiratory tract infections due to a human metapneumovirus in children: descriptive study and comparison with respiratory syncytial virus infections. Arch. Pediatr. 11:212–215. 11. Balada-Llasat, J. M., H. LaRue, C. Kelly, L. Rigali, and P. Pancholi. 2011. Evaluation of commercial ResPlex II v2.0, MultiCode-PLx, and xTAG respiratory viral panels for the diagnosis of respiratory viral infections in adults. J. Clin. Virol. 50:42–45. 12. Bao, X., et al. 2008. Human metapneumovirus small hydrophobic protein inhibits NF-kappaB transcriptional activity. J. Virol. 82:8224–8229. 13. Bao, X., et al. 2008. Human metapneumovirus glycoprotein G inhibits innate immune responses. PLoS Pathog. 4:e1000077. 14. Bao, X., et al. 2007. Airway epithelial cell response to human metapneumovirus infection. Virology 368:91–101. 15. Bao, X., et al. 2008. Identification of human metapneumovirus-induced gene networks in airway epithelial cells by microarray analysis. Virology 374:114–127. 16. Reference deleted. 17. Bastien, N., et al. 2003. Human metapneumovirus infection in the Canadian population. J. Clin. Microbiol. 41:4642–4646. 18. Ba ¨yon-Auboyer, M. H., C. T. D. Arnauld, and N. Eterradossi. 2000. Nucleotide sequences of the F, L and G protein genes of two non-A/non-B avian pneumoviruses (APV) reveal a novel APV subgroup. J. Gen. Virol. 81: 2723–2733. 19. Beneri, C., C. C. Ginocchio, R. Manji, and S. Sood. 2009. Comparison of clinical features of pediatric respiratory syncytial virus and human metapneumovirus infections. Infect. Control Hosp. Epidemiol. 30:1240–1241. 20. Bermingham, A., and P. L. Collins. 1999. The M2-2 protein of human respiratory syncytial virus is a regulatory factor involved in the balance between RNA replication and transcription. Proc. Natl. Acad. Sci. U. S. A. 96:11259–11264. 21. Bharaj, P., et al. 2009. Respiratory viral infections detected by multiplex PCR among pediatric patients with lower respiratory tract infections seen at an urban hospital in Delhi from 2005 to 2007. Virol. J. 6:89. 22. Biacchesi, S., et al. 2005. Infection of nonhuman primates with recombinant human metapneumovirus lacking the SH, G, or M2-2 protein categorizes each as a nonessential accessory protein and identifies vaccine candidates. J. Virol. 79:12608–12613. 23. Biacchesi, S., et al. 2006. Modification of the trypsin-dependent cleavage activation site of the human metapneumovirus fusion protein to be trypsin independent does not increase replication or spread in rodents or nonhuman primates. J. Virol. 80:5798–5806. 24. Biacchesi, S., et al. 2004. Recovery of human metapneumovirus from cDNA: optimization of growth in vitro and expression of additional genes. Virology 321:247–259. 25. Biacchesi, S., et al. 2004. Recombinant human metapneumovirus lacking the small hydrophobic SH and/or attachment G glycoprotein: deletion of G yields a promising vaccine candidate. J. Virol. 78:12877–12887. 26. Blanco, J. C., M. S. Boukhvalova, K. A. Shirey, G. A. Prince, and S. N. Vogel. New insights for development of a safe and protective RSV vaccine. Hum. Vaccin. 6:482–492. 27. Boeckh, M., V. Erard, D. Zerr, and J. Englund. 2005. Emerging viral infections after hematopoietic cell transplantation. Pediatr. Transplant. 9(Suppl. 7):48–54. 28. Boivin, G., et al. 2002. Virological features and clinical manifestations associated with human metapneumovirus: a new paramyxovirus responsible for acute respiratory-tract infections in all age groups. J. Infect. Dis. 186: 1330–1334. 29. Boivin, G., et al. 2003. Human metapneumovirus infections in hospitalized children. Emerg. Infect. Dis. 9:634–640. 30. Boivin, G., et al. 2007. An outbreak of severe respiratory tract infection due to human metapneumovirus in a long-term care facility. Clin. Infect. Dis. 44:1152–1158. 31. Bosis, S., et al. 2005. Impact of human metapneumovirus in childhood: comparison with respiratory syncytial virus and influenza viruses. J. Med. Virol. 75:101–104. 32. Boukhvalova, M. S., G. A. Prince, and J. C. Blanco. 2009. The cotton rat model of respiratory viral infections. Biologicals 37:152–159. 33. Brien, J., J. L. Uhrlaub, A. Hirsch, C. A. Wiley, and J. Nikolich-Zugich.

CLIN. MICROBIOL. REV.

34.

35.

36.

37.

38.

39.

40.

41. 42.

43.

44.

45.

46.

47. 48. 49. 50.

51.

52.

53.

54.

55. 56.

57. 58.

59.

60.

61.

2009. Key role of T cell defects in age-related vulnerability to West Nile virus. J. Exp. Med. 206:2735–2745. Bruijnesteijn van Coppenraet, L. E., et al. 2010. Comparison of two commercial molecular assays for simultaneous detection of respiratory viruses in clinical samples using two automatic electrophoresis detection systems. J. Virol. Methods 169:188–192. Buchholz, U. J., et al. 2005. Deletion of M2 gene open reading frames 1 and 2 of human metapneumovirus: effects on RNA synthesis, attenuation, and immunogenicity. J. Virol. 79:6588–6597. Burton, R. E. W., et al. 2010. A microfluid chip-compatible bioassay based on single-molecule detection with high sensitivity and multiplexing. Lab Chip 10:843–851. Byus, S. B., et al. 1989. Swollen head syndrome in chicken: a preliminary report on the isolation of a possible aetiological agent. J. S. Afr. Vet. Assoc. 60:221–222. Cane, P. A., B. G. van den Hoogen, S. Chakrabarti, C. D. Fegan, and A. D. Osterhaus. 2003. Human metapneumovirus in a haematopoietic stem cell transplant recipient with fatal lower respiratory tract disease. Bone Marrow Transplant. 31:309–310. Caracciolo, S., et al. 2008. Human metapneumovirus infection in young children hospitalized with acute respiratory tract disease: virologic and clinical features. Pediatr. Infect. Dis. J. 27:406–412. Carneiro, B. M., et al. 2009. Detection of all four human metapneumovirus subtypes in nasopharyngeal specimens from children with respiratory disease in Uberlandia, Brazil. J. Med. Virol. 81:1814–1818. Carr, M. J., et al. 2008. Molecular epidemiology of human metapneumovirus in Ireland. J. Med. Virol. 80:510–516. Chen, K. F., et al. 2011. Reverse transcription polymerase chain reaction and electrospray ionization mass spectrometry for identifying acute viral upper respiratory tract infections. Diagn. Microbiol. Infect. Dis. 69:179–186. Chiu, C. Y., et al. 2007. Diagnosis of a critical respiratory illness caused by human metapneumovirus by use of a pan-virus microarray. J. Clin. Microbiol. 45:2340–2343. Chung, J. Y., T. H. Han, S. W. Kim, and E. S. Hwang. 2008. Genotype variability of human metapneumovirus, South Korea. J. Med. Virol. 80: 902–905. Chung, J. Y., T. H. Han, S. W. Kim, and E. S. Hwang. 2007. Respiratory picornavirus infections in Korean children with lower respiratory tract infections. Scand. J. Infect. Dis. 39:250–254. Collins, P. L., M. G. Hill, J. Cristina, and H. Grosfeld. 1996. Transcription elongation factor of respiratory syncytial virus, a nonsegmented negativestrand RNA virus. Proc. Natl. Acad. Sci. U. S. A. 93:81–85. Cseke, G., et al. 2009. Integrin alphavbeta1 promotes infection by human metapneumovirus. Proc. Natl. Acad. Sci. U. S. A. 106:1566–1571. Cseke, G., et al. 2007. Human metapneumovirus fusion protein vaccines that are immunogenic and protective in cotton rats. J. Virol. 81:698–707. Cuevas, L. E., et al. 2003. Human metapneumovirus and respiratory syncytial virus, Brazil. Emerg. Infect. Dis. 9:1626–1628. Darniot, M., T. Petrella, S. Aho, P. Pothier, and C. Manoha. 2005. Immune response and alteration of pulmonary function after primary human metapneumovirus (hMPV) infection of BALB/c mice. Vaccine 23:4473–4480. Darniot, M., et al. 2009. Age-associated aggravation of clinical disease after primary metapneumovirus infection of BALB/c mice. J. Virol. 83:3323– 3332. David, P., D. Hirschwerk, S. Goldberg, and C. Ginocchio. 2009. Acute pericarditis caused by human metapneumovirus. Infect. Dis. Clin. Pract. 17:283–285. Debiaggi, M., et al. 2006. Persistent symptomless human metapneumovirus infection in hematopoietic stem cell transplant recipients. J. Infect. Dis. 194:474–478. de Graaf, M., et al. 2008. Specificity and functional interaction of the polymerase complex proteins of human and avian metapneumoviruses. J. Gen. Virol. 89:975–983. de Graaf, M., et al. 2007. An improved plaque reduction virus neutralization assay for human metapneumovirus. J. Virol. Methods 143:169–174. de Graaf, M., A. D. Osterhaus, R. A. Fouchier, and E. C. Holmes. 2008. Evolutionary dynamics of human and avian metapneumoviruses. J. Gen. Virol. 89:2933–2942. de Graaf, M., et al. 2009. Fusion protein is the main determinant of metapneumovirus host tropism. J. Gen. Virol. 90:1408–1416. de Graaf, M. H. S., et al. 2009. Metapneumovirus; determinants of host range and replication. Erasmus University Rotterdam, Rotterdam, The Netherlands. de Swart, R. L., et al. 2007. Immunization of macaques with formalininactivated human metapneumovirus induces hypersensitivity to hMPV infection. Vaccine 25:8518–8528. DeVincenzo, J. P., R. L. Hirsch, R. J. Fuentes, and F. H. Top, Jr. 2000. Respiratory syncytial virus immune globulin treatment of lower respiratory tract infection in pediatric patients undergoing bone marrow transplantation—a compassionate use experience. Bone Marrow Transplant. 25:161–165. Dinwiddie, D. L., and K. S. Harrod. 2008. Human metapneumovirus inhib-

VOL. 24, 2011

62.

63.

64.

65.

66. 67. 68.

69.

70. 71.

72. 73. 74.

75.

76.

77.

78.

79. 80.

81. 82.

83. 84. 85. 86.

87. 88.

89. 90.

91.

its IFN-alpha signaling through inhibition of STAT1 phosphorylation. Am. J. Respir. Cell Mol. Biol. 38:661–670. Ditt, V., J. Lusebrink, R. L. Tillmann, V. Schildgen, and O. Schildgen. 2011. Respiratory infections by HMPV and RSV are clinically indistinguishable but induce different host response in aged individuals. PLoS One 6:e16314. Dollner, H., K. Risnes, A. Radtke, and S. A. Nordbo. 2004. Outbreak of human metapneumovirus infection in Norwegian children. Pediatr. Infect. Dis. J. 23:436–440. Don, M., M. Korppi, F. Valent, R. Vainionpaa, and M. Canciani. 2008. Human metapneumovirus pneumonia in children: results of an Italian study and mini-review. Scand. J. Infect. Dis. 40:821–826. Drews, S. J., et al. 2008. Use of the Seeplex RV detection kit for surveillance of respiratory viral outbreaks in Toronto, Ontario, Canada. Ann. Clin. Lab. Sci. 38:376–379. Ebihara, T., et al. 2004. Early reinfection with human metapneumovirus in an infant. J. Clin. Microbiol. 42:5944–5946. Ebihara, T., et al. 2004. Human metapneumovirus infection in Japanese children. J. Clin. Microbiol. 42:126–132. Ebihara, T., et al. 2004. Comparison of the seroprevalence of human metapneumovirus and human respiratory syncytial virus. J. Med. Virol. 72:304–306. Englund, J. A., et al. 2006. Brief communication: fatal human metapneumovirus infection in stem-cell transplant recipients. Ann. Intern. Med. 144:344–349. Escobar, C., et al. 2009. Genetic variability of human metapneumovirus isolated from Chilean children, 2003–2004. J. Med. Virol. 81:340–344. Esper, F., D. Boucher, C. Weibel, R. A. Martinello, and J. S. Kahn. 2003. Human metapneumovirus infection in the United States: clinical manifestations associated with a newly emerging respiratory infection in children. Pediatrics 111:1407–1410. Esper, F., et al. 2004. A 1-year experience with human metapneumovirus in children aged ⬍5 years. J. Infect. Dis. 189:1388–1396. Fabbiani, M., et al. 2009. Epidemiological and clinical study of viral respiratory tract infections in children from Italy. J. Med. Virol. 81:750–756. Falsey, A. R., M. C. Criddle, and E. E. Walsh. 2006. Detection of respiratory syncytial virus and human metapneumovirus by reverse transcription polymerase chain reaction in adults with and without respiratory illness. J. Clin. Virol. 35:46–50. Falsey, A. R., D. Erdman, L. J. Anderson, and E. E. Walsh. 2003. Human metapneumovirus infections in young and elderly adults. J. Infect. Dis. 187:785–790. Falsey, A. R., M. A. Formica, J. J. Treanor, and E. E. Walsh. 2003. Comparison of quantitative reverse transcription-PCR to viral culture for assessment of respiratory syncytial virus shedding. J. Clin. Microbiol. 41:4160–4165. Falsey, A. R., M. A. Formica, and E. E. Walsh. 2009. Microneutralization assay for the measurement of neutralizing antibodies to human metapneumovirus. J. Clin. Virol. 46:314–317. Fearns, R., and P. L. Collins. 1999. Role of the M2-1 transcription antitermination protein of respiratory syncytial virus in sequential transcription. J. Virol. 73:5852–5864. Reference deleted. Franquet, T., et al. 2005. Human metapneumovirus infection in hematopoietic stem cell transplant recipients: high-resolution computed tomography findings. J. Comput. Assist. Tomogr. 29:223–227. Freymouth, F., et al. 2003. Presence of the new human metapneumovirus in French children with bronchiolitis. Pediatr. Infect. Dis. J. 22:92–94. Fuenzalida, L., et al. 2010. Usefulness of two new methods for diagnosing metapneumovirus infections in children. Clin. Microbiol. Infect. 16:1663– 1668. Furuse, Y., et al. Detection of novel respiratory viruses from influenza-like illness in the Philippines. J. Med. Virol. 82:1071–1074. Galiano, M., et al. 2004. Evidence of human metapneumovirus in children in Argentina. J. Med. Virol. 72:299–303. Garbino, J., et al. 2008. Respiratory viruses in HIV-infected patients with suspected respiratory opportunistic infection. AIDS 22:701–705. Garcia-Garcia, M. L., et al. 2007. Human metapneumovirus bronchiolitis in infancy is an important risk factor for asthma at age 5. Pediatr. Pulmonol. 42:458–464. Garcia-Garcia, M. L., et al. 2006. Human metapneumovirus infections in hospitalised infants in Spain. Arch. Dis. Child. 91:290–295. Garcia-Garcia, M. L., et al. 2006. Prevalence and clinical characteristics of human metapneumovirus infections in hospitalized infants in Spain. Pediatr. Pulmonol. 41:863–871. Garcia-Sastre, A. 2006. Antiviral response in pandemic influenza viruses. Emerg. Infect. Dis. 12:44–47. Gaunt, E., E. C. McWilliam-Leitch, K. Templeton, and P. Simmonds. 2009. Incidence, molecular epidemiology and clinical presentations of human metapneumovirus; assessment of its importance as a diagnostic screening target. J. Clin. Virol. 46:318–324. Gerna, G., et al. 2005. Changing circulation rate of human metapneumovirus strains and types among hospitalized pediatric patients during three consecutive winter-spring seasons. Brief report. Arch. Virol. 150:2365–2375.

HUMAN METAPNEUMOVIRUS

749

92. Ginocchio, C. C., R. Manji, M. Lotlikar, and F. Zhang. 2008. Clinical evaluation of NucliSENS magnetic extraction and NucliSENS analyte-specific reagents for real-time detection of human metapneumovirus in pediatric respiratory specimens. J. Clin. Microbiol. 46:1274–1280. 93. Gioula, G., D. Chatzidimitriou, A. Melidou, M. Exindari, and V. Kyriazopoulou-Dalaina. 2010. Contribution of human metapneumovirus to influenza-like infections in North Greece, 2005–2008. Euro. Surveill. 15:19499. 94. Gould, P. S., and A. J. Easton. 2007. Coupled translation of the second open reading frame of M2 mRNA is sequence dependent and differs significantly within the subfamily Pneumovirinae. J. Virol. 81:8488–8496. 95. Goutagny, N., et al. 2010. Cell type-specific recognition of human metapneumoviruses (HMPVs) by retinoic acid-inducible gene I (RIG-I) and TLR7 and viral interference of RIG-I ligand recognition by HMPV-B1 phosphoprotein. J. Immunol. 184:1168–1179. 96. Gray, G. C., et al. 2006. Multi-year study of human metapneumovirus infection at a large US midwestern medical referral center. J. Clin. Virol. 37:269–276. 97. Gray, G. C., et al. 2006. Human metapneumovirus, Peru. Emerg. Infect. Dis. 12:347–350. 98. Greensill, J., et al. 2003. Human metapneumovirus in severe respiratory syncytial virus bronchiolitis. Emerg. Infect. Dis. 9:372–375. 99. Guerrero-Plata, A., et al. 2005. Activity and regulation of alpha interferon in respiratory syncytial virus and human metapneumovirus experimental infections. J. Virol. 79:10190–10199. 100. Guerrero-Plata, A., et al. 2006. Differential response of dendritic cells to human metapneumovirus and respiratory syncytial virus. Am. J. Respir. Cell Mol. Biol. 34:320–329. 101. Guerrero-Plata, A., D. Kolli, C. Hong, A. Casola, and R. P. Garofalo. 2009. Subversion of pulmonary dendritic cell function by paramyxovirus infections. J. Immunol. 182:3072–3083. 102. Hamelin, M. E., Y. Abed, and G. Boivin. 2004. Human metapneumovirus: a new player among respiratory viruses. Clin. Infect. Dis. 38:983–990. 103. Hamelin, M. E., et al. 2005. Human metapneumovirus infection in adults with community-acquired pneumonia and exacerbation of chronic obstructive pulmonary disease. Clin. Infect. Dis. 41:498–502. 104. Hamelin, M. E., G. A. Prince, and G. Boivin. 2006. Effect of ribavirin and glucocorticoid treatment in a mouse model of human metapneumovirus infection. Antimicrob. Agents Chemother. 50:774–777. 105. Hamelin, M. E., G. A. Prince, A. M. Gomez, R. Kinkead, and G. Boivin. 2006. Human metapneumovirus infection induces long-term pulmonary inflammation associated with airway obstruction and hyperresponsiveness in mice. J. Infect. Dis. 193:1634–1642. 106. Hamelin, M. E., et al. 2005. Pathogenesis of human metapneumovirus lung infection in BALB/c mice and cotton rats. J. Virol. 79:8894–8903. 107. Heininger, U., A. T. Kruker, J. Bonhoeffer, and U. B. Schaad. 2009. Human metapneumovirus infections—biannual epidemics and clinical findings in children in the region of Basel, Switzerland. Eur. J. Pediatr. 168:1455–1460. 108. Herd, K. A., et al. 2006. Cytotoxic T-lymphocyte epitope vaccination protects against human metapneumovirus infection and disease in mice. J. Virol. 80:2034–2044. 109. Herd, K. A., M. D. Nissen, P. M. Hopkins, T. P. Sloots, and R. W. Tindle. 2008. Major histocompatibility complex class I cytotoxic T lymphocyte immunity to human metapneumovirus (hMPV) in individuals with previous hMPV infection and respiratory disease. J. Infect. Dis. 197:584–592. 110. Herfst, S., et al. 2004. Recovery of human metapneumovirus genetic lineages A and B from cloned cDNA. J. Virol. 78:8264–8270. 111. Herfst, S., et al. 2008. Generation of temperature-sensitive human metapneumovirus strains that provide protective immunity in hamsters. J. Gen. Virol. 89:1553–1562. 112. Herfst, S., et al. 2007. Immunization of Syrian golden hamsters with F subunit vaccine of human metapneumovirus induces protection against challenge with homologous or heterologous strains. J. Gen. Virol. 88:2702–2709. 113. Herfst, S., and R. A. Fouchier. 2008. Vaccination approaches to combat human metapneumovirus lower respiratory tract infections. J. Clin. Virol. 41:49–52. 114. Herfst, S., et al. 2008. Low-pH-induced membrane fusion mediated by human metapneumovirus F protein is a rare, strain-dependent phenomenon. J. Virol. 82:8891–8895. 115. High, K. P. 2005. Pneumonia in older adults. New categories add complexity to diagnosis and care. Postgrad. Med. 118:18–20, 25–28. 116. Honda, H., et al. 2006. Outbreak of human metapneumovirus infection in elderly inpatients in Japan. J. Am. Geriatr. Soc. 54:177–180. 117. Hopkins, P., et al. 2008. Human metapneumovirus in lung transplant recipients and comparison to respiratory syncitial virus. Am. J. Respir. Crit. Care Med. 178:876–881. 118. Huck, B., et al. 2006. Novel human metapneumovirus sublineage. Emerg. Infect. Dis. 12:147–150. 118a.IJpma, F. F., et al. 2004. Human metapneumovirus infection in hospital referred South African children. J. Med. Virol. 73:486–493. 119. Jartti, T., B. van den Hoogen, R. P. Garofalo, A. D. Osterhaus, and O. Ruuskanen. 2002. Metapneumovirus and acute wheezing in children. Lancet 360:1393–1394.

750

SCHILDGEN ET AL.

120. Jenkins, G. M. R., et al. 2002. Rates of molecular evolution in RNA viruses: a quantitative phylogenetic analysis. J. Mol. Evol. 54:156–165. 121. Jiang, J., et al. 2009. Limited expansion of virus-specific CD8 T cells in the aged environment. Mech. Ageing Dev. 130:713–721. 122. Jiang, T., et al. 2009. CpG oligodeoxynucleotides protect against the H1N1 pandemic influenza virus infection in a murine model. Antiviral Res. 89: 124–126. 123. Jiang, T. J., et al. 2009. Preferential loss of Th17 cells is associated with CD4 T cell activation in patients with pandemic H1N1 swine-origin influenza A infection. Clin. Immunol. 137:303–310. 124. Jokela, P., H. Piiparinen, K. Luiro, and M. Lappalainen. 2010. Detection of human metapneumovirus and respiratory syncytial virus by duplex realtime RT-PCR assay in comparison with direct fluorescent assay. Clin. Microbiol. Infect. 16:1568–1573. 125. Kahn, J. S., M. J. Schnell, L. Buonocore, and J. K. Rose. 1999. Recombinant vesicular stomatitis virus expressing respiratory syncytial virus (RSV) glycoproteins: RSV fusion protein can mediate infection and cell fusion. Virology 254:81–91. 126. Kaida, A., et al. 2006. Seasonal distribution and phylogenetic analysis of human metapneumovirus among children in Osaka City, Japan. J. Clin. Virol. 35:394–399. 127. Kawai, T. A. S. 2005. Pathogen recognotion with Toll-like receptors. Curr. Opin. Immunol. 17:338–344. 128. Kaye, M., S. Skidmore, H. Osman, M. Weinbren, and R. Warren. 2006. Surveillance of respiratory virus infections in adult hospital admissions using rapid methods. Epidemiol. Infect. 134:792–798. 129. Kolli, D., et al. 2008. T lymphocytes contribute to antiviral immunity and pathogenesis in experimental human metapneumovirus infection. J. Virol. 82:8560–8569. 130. Konig, B., et al. 2004. Prospective study of human metapneumovirus infection in children less than 3 years of age. J. Clin. Microbiol. 42:4632–4635. 131. Kuiken, T., et al. 2004. Experimental human metapneumovirus infection of cynomolgus macaques (Macaca fascicularis) results in virus replication in ciliated epithelial cells and pneumocytes with associated lesions throughout the respiratory tract. Am. J. Pathol. 164:1893–1900. 132. Laguna-Torres, V. A., et al. 2011. Influenza and other respiratory viruses in three Central American countries. Influenza Other Respi. Viruses 5:123– 134. 133. Laham, F. R., et al. 2004. Differential production of inflammatory cytokines in primary infection with human metapneumovirus and with other common respiratory viruses of infancy. J. Infect. Dis. 189:2047–2056. 134. Lamb, R. A. 1993. Paramyxovirus fusion: a hypothesis for changes. Virology 197:1–11. 135. Lamb, R. A., and T. S. Jardetzky. 2007. Structural basis of viral invasion: lessons from paramyxovirus F. Curr. Opin. Struct. Biol. 17:427–436. 136. Lamb, R. A. P., and G. D. Parks. 2007. Paramyxoviridae: the viruses and their replication, p. 1449–1456. In D. M. Knipe et al. (ed.), Fields virology, 5th ed. Lippincott Williams & Wilkins, Philadelphia, PA. 137. Landry, M. L., S. Cohen, and D. Ferguson. 2008. Prospective study of human metapneumovirus detection in clinical samples by use of Light Diagnostics direct immunofluorescence reagent and real-time PCR. J. Clin. Microbiol. 46:1098–1100. 138. Landry, M. L., D. Ferguson, S. Cohen, T. C. Peret, and D. D. Erdman. 2005. Detection of human metapneumovirus in clinical samples by immunofluorescence staining of shell vial centrifugation cultures prepared from three different cell lines. J. Clin. Microbiol. 43:1950–1952. 139. Larcher, C., et al. 2005. Human metapneumovirus infection in lung transplant recipients: clinical presentation and epidemiology. J. Heart Lung Transplant. 24:1891–1901. 140. Larcher, C., et al. 2008. Comparison of human metapneumovirus genotypes from the province of Bolzano in northern Italy with strains from surrounding regions in Italy and Austria. Jpn. J. Infect. Dis. 61:154–156. 141. Lazar, I., et al. 2004. Human metapneumovirus and severity of respiratory syncytial virus disease. Emerg. Infect. Dis. 10:1318–1320. 142. Leung, J., F. Esper, C. Weibel, and J. S. Kahn. 2005. Seroepidemiology of human metapneumovirus (hMPV) on the basis of a novel enzyme-linked immunosorbent assay utilizing hMPV fusion protein expressed in recombinant vesicular stomatitis virus. J. Clin. Microbiol. 43:1213–1219. 143. Liao, S., et al. 2008. Role of retinoic acid inducible gene-I in human metapneumovirus-induced cellular signalling. J. Gen. Virol. 89:1978–1986. 144. Liu, L., N. Bastien, and Y. Li. 2007. Intracellular processing, glycosylation, and cell surface expression of human metapneumovirus attachment glycoprotein. J. Virol. 81:13435–13443. 145. Liu, L., N. Bastien, F. Sidaway, E. Chan, and Y. Li. 2007. Seroprevalence of human metapneumovirus (hMPV) in the Canadian province of Saskatchewan analyzed by a recombinant nucleocapsid protein-based enzyme-linked immunosorbent assay. J. Med. Virol. 79:308–313. 146. Ljubin-Sternak, S., et al. 2008. Detection of genetic lineages of human metapneumovirus in Croatia during the winter season 2005/2006. J. Med. Virol. 80:1282–1287. 147. Ljubin Sternak, S., T. Vilibic Cavlek, A. R. Falsey, E. E. Walsh, and G.

CLIN. MICROBIOL. REV.

148. 149. 150.

151.

152. 153.

154. 155.

156.

157.

158.

159.

160.

161.

162.

163. 164.

165. 166.

167.

168. 169.

170.

171.

172. 173.

174.

175. 176.

Mlinaric Galinovic. 2006. Serosurvey of human metapneumovirus infection in Croatia. Croat. Med. J. 47:878–881. Loo, L. H., et al. 2007. Human metapneumovirus in children, Singapore. Emerg. Infect. Dis. 13:1396–1398. Louie, J. K., et al. 2007. A summer outbreak of human metapneumovirus infection in a long-term-care facility. J. Infect. Dis. 196:705–708. Lu, G., et al. Large-scale seroprevalence analysis of human metapneumovirus and human respiratory syncytial virus infections in Beijing, China. Virol. J. 8:62. Luchsinger, V., C. Escobar, and L. F. Avendano. 2005. Detection of human metapneumovirus in children hospitalized for acute lower respiratory infection in Santiago, Chile. Rev. Med. Chil. 133:1059–1064. Ludewick, H. P., et al. 2005. Human metapneumovirus genetic variability, South Africa. Emerg. Infect. Dis. 11:1074–1078. Lusebrink, J., et al. High seroprevalence of neutralizing capacity against human metapneumovirus in all age groups studied in Bonn, Germany. Clin. Vaccine Immunol. 17:481–484. Mackay, I. M., et al. 2006. Genetic diversity of human metapneumovirus over 4 consecutive years in Australia. J. Infect. Dis. 193:1630–1633. MacPhail, M., et al. 2004. Identification of small-animal and primate models for evaluation of vaccine candidates for human metapneumovirus (hMPV) and implications for hMPV vaccine design. J. Gen. Virol. 85:1655– 1663. Madhi, S. A., H. Ludewick, Y. Abed, K. P. Klugman, and G. Boivin. 2003. Human metapneumovirus-associated lower respiratory tract infections among hospitalized human immunodeficiency virus type 1 (HIV-1)-infected and HIV-1-uninfected African infants. Clin. Infect. Dis. 37:1705–1710. Madhi, S. A., et al. 2007. Seasonality, incidence, and repeat human metapneumovirus lower respiratory tract infections in an area with a high prevalence of human immunodeficiency virus type-1 infection. Pediatr. Infect. Dis. J. 26:693–699. Maertzdorf, J., et al. 2004. Real-time reverse transcriptase PCR assay for detection of human metapneumoviruses from all known genetic lineages. J. Clin. Microbiol. 42:981–986. Maggi, F., et al. 2003. Human metapneumovirus associated with respiratory tract infections in a 3-year study of nasal swabs from infants in Italy. J. Clin. Microbiol. 41:2987–2991. Manoha, C., S. Espinosa, S. L. Aho, F. Huet, and P. Pothier. 2007. Epidemiological and clinical features of hMPV, RSV and RVs infections in young children. J. Clin. Virol. 38:221–226. Mao, H. W., X. Q. Yang, and X. D. Zhao. 2008. Characterization of human metapneumoviruses isolated in Chongqing, China. Chin. Med. J. (Engl.) 121:2254–2257. Martinello, R. A., M. D. Chen, C. Weibel, and J. S. Kahn. 2002. Correlation between respiratory syncytial virus genotype and severity of illness. J. Infect. Dis. 186:839–842. Martinello, R. A., et al. 2006. Human metapneumovirus and exacerbations of chronic obstructive pulmonary disease. J. Infect. 53:248–254. Martino, R., et al. 2005. Prospective study of the incidence, clinical features, and outcome of symptomatic upper and lower respiratory tract infections by respiratory viruses in adult recipients of hematopoietic stem cell transplants for hematologic malignancies. Biol. Blood Marrow Transplant. 11:781–796. Mathisen, M., et al. 2009. RNA viruses in community-acquired childhood pneumonia in semi-urban Nepal; a cross-sectional study. BMC Med. 7:35. Matsuzaki, Y., et al. 2008. Clinical impact of human metapneumovirus genotypes and genotype-specific seroprevalence in Yamagata, Japan. J. Med. Virol. 80:1084–1089. Mazhul, L. A., O. Schildgen, E. I. Isaeva, and S. O. Viazov. 2007. Human metapneumovirus as a common cause of respiratory tract disease. Vopr. Virusol. 52:4–8. McAdam, A. J., et al. 2004. Human metapneumovirus in children tested at a tertiary-care hospital. J. Infect. Dis. 190:20–26. Melendi, G. A., et al. 2007. Mapping and characterization of the primary and anamnestic H-2(d)-restricted cytotoxic T-lymphocyte response in mice against human metapneumovirus. J. Virol. 81:11461–11467. Mirazo, S., D. Ruchansky, A. Blanc, and J. Arbiza. 2005. Serologic evidence of human metapneumovirus circulation in Uruguay. Mem. Inst. Oswaldo Cruz 100:715–718. Mok, H., et al. 2008. An alphavirus replicon-based human metapneumovirus vaccine is immunogenic and protective in mice and cotton rats. J. Virol. 82:11410–11418. Monto, A. S. 2002. Epidemiology of viral respiratory infections. Am. J. Med. 112(Suppl. 6A):4S–12S. Mullins, J. A., et al. 2004. Human metapneumovirus infection among children hospitalized with acute respiratory illness. Emerg. Infect. Dis. 10:700– 705. Nicholson, K. G., T. McNally, M. Silverman, P. Simons, and M. C. Zambon. 2003. Influenza-related hospitalizations among young children in Leicestershire. Pediatr. Infect. Dis. J. 22:S228–S230. Nolte, F. S., et al. 2007. MultiCode-PLx system for multiplexed detection of seventeen respiratory viruses. J. Clin. Microbiol. 45:2779–2786. Okamoto, M., et al. 2010. Development and evaluation of a whole virus-

VOL. 24, 2011

177.

178.

179. 180.

181.

182. 183.

184.

185.

186. 187.

188.

189.

190.

191.

192. 193.

194. 195. 196.

197.

198.

199.

200.

201. 202.

203.

based enzyme-linked immunosorbent assay for the detection of human metapneumovirus antibodies in human sera. J. Virol. Methods 164:24–29. Oliveira, D. B., et al. 2009. Epidemiology and genetic variability of human metapneumovirus during a 4-year-long study in Southeastern Brazil. J. Med. Virol. 81:915–921. Olmsted, R. A., et al. 1986. Expression of the F glycoprotein of respiratory syncytial virus by a recombinant vaccinia virus: comparison of the individual contributions of the F and G glycoproteins to host immunity. Proc. Natl. Acad. Sci. U. S. A. 83:7462–7466. Ordas, J., et al. 2006. Role of metapneumovirus in viral respiratory infections in young children. J. Clin. Microbiol. 44:2739–2742. Osbourn, M., K. A. McPhie, V. M. Ratnamohan, D. E. Dwyer, and D. N. Durrheim. 2009. Outbreak of human metapneumovirus infection in a residential aged care facility. Commun. Dis. Intell. 33:38–40. Pavlin, J. A., et al. 2008. Human metapneumovirus reinfection among children in Thailand determined by ELISA using purified soluble fusion protein. J. Infect. Dis. 198:836–842. Peiris, J. S., et al. 2003. Children with respiratory disease associated with metapneumovirus in Hong Kong. Emerg. Infect. Dis. 9:628–633. Pelletier, G., P. Dery, Y. Abed, and G. Boivin. 2002. Respiratory tract reinfections by the new human metapneumovirus in an immunocompromised child. Emerg. Infect. Dis. 8:976–978. Pham, Q. N., et al. 2005. Chimeric recombinant human metapneumoviruses with the nucleoprotein or phosphoprotein open reading frame replaced by that of avian metapneumovirus exhibit improved growth in vitro and attenuation in vivo. J. Virol. 79:15114–15122. Pitoiset, C., et al. 2010. Human metapneumovirus genotypes and severity of disease in young children (n ⫽ 100) during a 7-year study in Dijon hospital, France. J. Med. Virol. 82:1782–1789. Pizzorno, A., et al. Molecular detection and genetic variability of human metapneumovirus in Uruguay. J. Med. Virol. 82:861–865. Po, J. L., E. M. Gardner, F. Anaraki, P. D. Katsikis, and D. Murasko. 2002. Age-associated decrease in virus-specific CD8⫹ T lymphocytes during primary influenza infection. Mech. Ageing Dev. 123:1167–1181. Rafiefard, F., Z. Yun, and C. Orvell. 2008. Epidemiologic characteristics and seasonal distribution of human metapneumovirus infections in five epidemic seasons in Stockholm, Sweden, 2002–2006. J. Med. Virol. 80: 1631–1638. Rawlinson, W. D., et al. 2003. Asthma exacerbations in children associated with rhinovirus but not human metapneumovirus infection. J. Infect. Dis. 187:1314–1318. Raymond, F., et al. 2009. Comparison of automated microarray detection with real-time PCR assays for detection of respiratory viruses in specimens obtained from children. J. Clin. Microbiol. 47:743–750. Raza, K., et al. 2007. Successful outcome of human metapneumovirus (hMPV) pneumonia in a lung transplant recipient treated with intravenous ribavirin. J. Heart Lung Transplant. 26:862–864. Reijans, M., et al. 2008. RespiFinder: a new multiparameter test to differentially identify fifteen respiratory viruses. J. Clin. Microbiol. 46:1232–1240. Robinson, J. L., B. E. Lee, N. Bastien, and Y. Li. 2005. Seasonality and clinical features of human metapneumovirus infection in children in Northern Alberta. J. Med. Virol. 76:98–105. Rohde, G., et al. 2005. Relevance of human metapneumovirus in exacerbations of COPD. Respir. Res. 6:150. Russell, C. J., and L. E. Luque. 2006. The structural basis of paramyxovirus invasion. Trends Microbiol. 14:243–246. Ryder, A. B., S. J. Tollefson, A. B. Podsiad, J. E. Johnson, and J. V. Williams. 2010. Soluble recombinant human metapneumovirus G protein is immunogenic but not protective. Vaccine 28:4145–4152. Schickli, J. H., et al. 2008. Deletion of human metapneumovirus M2-2 increases mutation frequency and attenuates growth in hamsters. Virol. J. 5:69. Schickli, J. H., J. Kaur, N. Ulbrandt, R. R. Spaete, and R. S. Tang. 2005. An S101P substitution in the putative cleavage motif of the human metapneumovirus fusion protein is a major determinant for trypsin-independent growth in Vero cells and does not alter tissue tropism in hamsters. J. Virol. 79:10678–10689. Schildgen, O., T. Geikowski, T. Glatzel, J. Schuster, and A. Simon. 2005. Frequency of human metapneumovirus in the upper respiratory tract of children with symptoms of an acute otitis media. Eur. J. Pediatr. 164:400– 401. Schildgen, O., et al. 2004. New variant of the human metapneumovirus (HMPV) associated with an acute and severe exacerbation of asthma bronchiale. J. Clin. Virol. 31:283–288. Schildgen, O., A. Simon, and J. Williams. 2007. Animal models for human metapneumovirus (HMPV) infections. Vet. Res. 38:117–126. Schildgen, V., et al. 2010. Human HepG2 cells support respiratory syncytial virus and human metapneumovirus replication. J. Virol. Methods 163:74–81. Schowalter, R. M., S. E. Smith, and R. E. Dutch. 2006. Characterization of human metapneumovirus F protein-promoted membrane fusion: critical roles for proteolytic processing and low pH. J. Virol. 80:10931–10941.

HUMAN METAPNEUMOVIRUS

751

204. Seal, B. S. S., et al. 2000. Fusion protein predicted amino acid sequence of the first US avian pneumovirus isolate and lack of heterogeneity among other US isolates. Virus Res. 66:139–147. 205. Semple, M. G., et al. 2005. Dual infection of infants by human metapneumovirus and human respiratory syncytial virus is strongly associated with severe bronchiolitis. J. Infect. Dis. 191:382–386. 206. Seth, R. B. S., et al. 2005. Identification and characterization of MAVS, a mitochondrial antiviral signalling protein that actrivates NF-kappa B and IRF-3. Cell 122:669–682. 207. Shanmukh, S., et al. 2008. Identification and classification of respiratory syncytial virus (RSV) strains by surface-enhanced Raman spectroscopy and multivariate statistical techniques. Anal. Bioanal. Chem. 390:1551–1555. 208. Singleton, R. J., et al. 2010. Viral respiratory infections in hospitalized and community control children in Alaska. J. Med. Virol. 82:1282–1290. 209. Skiadopoulos, M. H., et al. 2006. Individual contributions of the human metapneumovirus F, G, and SH surface glycoproteins to the induction of neutralizing antibodies and protective immunity. Virology 345:492–501. 210. Skiadopoulos, M. H., et al. 2004. The two major human metapneumovirus genetic lineages are highly related antigenically, and the fusion (F) protein is a major contributor to this antigenic relatedness. J. Virol. 78:6927–6937. 211. Sloots, T. P., et al. 2006. Human metapneumovirus, Australia, 2001–2004. Emerg. Infect. Dis. 12:1263–1266. 212. Smith, E. C., A. Popa, A. Chang, C. Masante, and R. E. Dutch. 2009. Viral entry mechanisms: the increasing diversity of paramyxovirus entry. FEBS J. 276:7217–7227. 213. Spetch, L., T. L. Bowlin, and A. Casola. 2008. Effect of NMSO3 treatment in a murine model of human metapneumovirus infection. J. Gen. Virol. 89:2709–2712. 214. Stout-Delgado, H. W., W. Du, A. C. Shirali, C. J. Booth, and D. Goldstein. 2009. Aging promotes neutrophil-induced mortality by augmenting IL-17 production during viral infection. Cell Host Microbe 6:446–456. 215. Sullender, W. M. 2000. Respiratory syncytial virus genetic and antigenic diversity. Clin. Microbiol. Rev. 13:1–15. 216. Sumino, K. C., et al. 2005. Detection of severe human metapneumovirus infection by real-time polymerase chain reaction and histopathological assessment. J. Infect. Dis. 192:1052–1060. 217. Suzuki, A., et al. 2005. Detection of human metapneumovirus from children with acute otitis media. Pediatr. Infect. Dis. J. 24:655–657. 218. Takaoka, A. Y. H. 2006. Interferon signalling network in innate defense. Cell. Microbiol. 8:907–922. 219. Talavera, G. A., and N. E. Mezquita. 2007. Human metapneumovirus in children with influenza-like illness in Yucatan, Mexico. Am. J. Trop. Med. Hyg. 76:182–183. 220. Tang, R. S., et al. 2004. Parainfluenza virus type 3 expressing the native or soluble fusion (F) protein of respiratory syncytial virus (RSV) confers protection from RSV infection in African green monkeys. J. Virol. 78: 11198–11207. 221. Tang, R. S., et al. 2005. A host-range restricted parainfluenza virus type 3 (PIV3) expressing the human metapneumovirus (hMPV) fusion protein elicits protective immunity in African green monkeys. Vaccine 23:1657–1667. 222. Tang, R. S., et al. 2003. Effects of human metapneumovirus and respiratory syncytial virus antigen insertion in two 3⬘ proximal genome positions of bovine/human parainfluenza virus type 3 on virus replication and immunogenicity. J. Virol. 77:10819–10828. 223. Tecu, C., et al. 2007. First detection of human metapneumovirus in children with respiratory infections in Romania. Roum. Arch. Microbiol. Immunol. 66:37–40. 224. Thammawat, S., T. A. Sadlon, P. G. Hallsworth, and D. L. Gordon. 2008. Role of cellular glycosaminoglycans and charged regions of viral G protein in human metapneumovirus infection. J. Virol. 82:11767–11774. 225. Thanasugarn, W., et al. 2003. Human metapneumovirus infection in Thai children. Scand. J. Infect. Dis. 35:754–756. 226. Tollefson, S. J., R. G. Cox, and J. V. Williams. 2010. Studies of culture conditions and environmental stability of human metapneumovirus. Virus Res. 151:54–59. 227. Ulbrandt, N. D., et al. 2006. Isolation and characterization of monoclonal antibodies which neutralize human metapneumovirus in vitro and in vivo. J. Virol. 80:7799–7806. 228. Ulloa-Gutierrez, R., et al. 2009. Human metapneumovirus in Costa Rica. Pediatr. Infect. Dis. J. 28:452–453. 229. van den Hoogen, B. G. 2007. Respiratory tract infection due to human metapneumovirus among elderly patients. Clin. Infect. Dis. 44:1159–1160. 230. van den Hoogen, B. G., T. M. Bestebroer, A. D. Osterhaus, and R. A. Fouchier. 2002. Analysis of the genomic sequence of a human metapneumovirus. Virology 295:119–132. 231. van den Hoogen, B. G., et al. 2001. A newly discovered human pneumovirus isolated from young children with respiratory tract disease. Nat. Med. 7:719–724. 232. van den Hoogen, B. G., et al. 2007. Experimental infection of macaques with human metapneumovirus induces transient protective immunity. J. Gen. Virol. 88:1251–1259.

752

SCHILDGEN ET AL.

CLIN. MICROBIOL. REV.

233. van den Hoogen, B. G., et al. 2004. Antigenic and genetic variability of human metapneumoviruses. Emerg. Infect. Dis. 10:658–666. 234. van den Hoogen, B. G., D. M. Osterhaus, and R. A. Fouchier. 2004. Clinical impact and diagnosis of human metapneumovirus infection. Pediatr. Infect. Dis. J. 23:S25–S32. 235. van den Hoogen, B. G., et al. 2003. Prevalence and clinical symptoms of human metapneumovirus infection in hospitalized patients. J. Infect. Dis. 188:1571–1577. 236. van Woensel, J. B., A. P. Bos, R. Lutter, J. W. Rossen, and R. Schuurman. 2006. Absence of human metapneumovirus co-infection in cases of severe respiratory syncytial virus infection. Pediatr. Pulmonol. 41:872–874. 237. Vargas, S. O., H. P. Kozakewich, A. R. Perez-Atayde, and A. J. McAdam. 2004. Pathology of human metapneumovirus infection: insights into the pathogenesis of a newly identified respiratory virus. Pediatr. Dev. Pathol. 7:478–486. 238. Viazov, S., F. Ratjen, R. Scheidhauer, M. Fiedler, and M. Roggendorf. 2003. High prevalence of human metapneumovirus infection in young children and genetic heterogeneity of the viral isolates. J. Clin. Microbiol. 41:3043–3045. 239. Vicente, D., M. Montes, G. Cilla, and E. Perez-Trallero. 2004. Human metapneumovirus and chronic obstructive pulmonary disease. Emerg. Infect. Dis. 10:1338–1339. 240. Vicente, D., M. Montes, G. Cilla, E. G. Perez-Yarza, and E. Perez-Trallero. 2006. Differences in clinical severity between genotype A and genotype B human metapneumovirus infection in children. Clin. Infect. Dis. 42:e111–e113. 241. Walsh, E. E., D. R. Peterson, and A. R. Falsey. 2008. Human metapneumovirus infections in adults: another piece of the puzzle. Arch. Intern. Med. 168:2489–2496. 242. Wang, H. C., et al. 2008. Co-circulating genetically divergent A2 human metapneumovirus strains among children in southern Taiwan. Arch. Virol. 153:2207–2213. 243. Wang, S. M., C. C. Liu, H. C. Wang, I. J. Su, and J. R. Wang. 2006. Human metapneumovirus infection among children in Taiwan: a comparison of clinical manifestations with other virus-associated respiratory tract infections. Clin. Microbiol. Infect. 12:1221–1224. 244. Weigl, J. A., et al. 2007. Ten years’ experience with year-round active surveillance of up to 19 respiratory pathogens in children. Eur. J. Pediatr. 166:957–966. 245. Wilkesmann, A., et al. 2006. Human metapneumovirus infections cause similar symptoms and clinical severity as respiratory syncytial virus infections. Eur. J. Pediatr. 165:467–475. 246. Williams, J. V., et al. 2007. A recombinant human monoclonal antibody to human metapneumovirus fusion protein that neutralizes virus in vitro and is effective therapeutically in vivo. J. Virol. 81:8315–8324. 247. Williams, J. V., et al. 2005. Human metapneumovirus infection plays an etiologic role in acute asthma exacerbations requiring hospitalization in adults. J. Infect. Dis. 192:1149–1153. 248. Williams, J. V., et al. 2004. Human metapneumovirus and lower respiratory tract disease in otherwise healthy infants and children. N. Engl. J. Med. 350:443–450.

249. Williams, J. V., et al. 2005. A prospective study comparing human metapneumovirus with other respiratory viruses in adults with hematologic malignancies and respiratory tract infections. J. Infect. Dis. 192:1061–1065. 250. Williams, J. V., et al. 2005. Human metapneumovirus infection in children hospitalized for wheezing. J. Allergy Clin. Immunol. 115:1311–1312. 251. Williams, J. V., S. J. Tollefson, J. E. Johnson, and J. E. Crowe, Jr. 2005. The cotton rat (Sigmodon hispidus) is a permissive small animal model of human metapneumovirus infection, pathogenesis, and protective immunity. J. Virol. 79:10944–10951. 252. Williams, J. V., S. J. Tollefson, S. Nair, and T. Chonmaitree. 2006. Association of human metapneumovirus with acute otitis media. Int. J. Pediatr. Otorhinolaryngol. 70:1189–1193. 253. Williams, J. V., et al. 2006. The role of human metapneumovirus in upper respiratory tract infections in children: a 20-year experience. J. Infect. Dis. 193:387–395. 254. Wolf, D. G., et al. 2006. Comparison of human metapneumovirus, respiratory syncytial virus and influenza A virus lower respiratory tract infections in hospitalized young children. Pediatr. Infect. Dis. J. 25:320–324. 255. Wolf, D. G., Z. Zakay-Rones, A. Fadeela, D. Greenberg, and R. Dagan. 2003. High seroprevalence of human metapneumovirus among young children in Israel. J. Infect. Dis. 188:1865–1867. 256. Wyde, P. R., S. N. Chetty, A. M. Jewell, G. Boivin, and P. A. Piedra. 2003. Comparison of the inhibition of human metapneumovirus and respiratory syncytial virus by ribavirin and immune serum globulin in vitro. Antiviral Res. 60:51–59. 257. Wyde, P. R., S. N. Chetty, A. M. Jewell, S. L. Schoonover, and P. A. Piedra. 2005. Development of a cotton rat-human metapneumovirus (hMPV) model for identifying and evaluating potential hMPV antivirals and vaccines. Antiviral Res. 66:57–66. 258. Xiao, N. G., et al. 2010. Prevalence and clinical and molecular characterization of human metapneumovirus in children with acute respiratory infection in China. Pediatr. Infect. Dis. J. 29:131–134. 259. Yang, C. F., et al. 2009. Genetic diversity and evolution of human metapneumovirus fusion protein over twenty years. Virol. J. 6:138. 260. Yim, K. C., et al. 2007. Human metapneumovirus: enhanced pulmonary disease in cotton rats immunized with formalin-inactivated virus vaccine and challenged. Vaccine 25:5034–5040. 261. Zeng, L., et al. 2011. Association between human metapneumovirus seroprevalence and hypertension in elderly subjects in a long-term care facility. Hypertens. Res. 34:474–478. 262. Zhang, Q., X. Q. Yang, Y. Zhao, and X. D. Zhao. 2008. High seroprevalence of human metapneumovirus infection in children in Chongqing, China. Chin. Med. J. (Engl.) 121:2162–2166. 263. Zlateva, K. T., P. Lemey, A. M. Vandamme, and M. Van Ranst. 2004. Molecular evolution and circulation patterns of human respiratory syncytial virus subgroup A: positively selected sites in the attachment G glycoprotein. J. Virol. 78:4675–4683.

Verena Schildgen studied biology at the University of Cologne. Her diploma thesis was on recombinant T cell receptors and their utility as therapeutic molecules. She performed her Ph.D. research at the University of Du ¨sseldorf and specialized in the molecular mechanisms of myelodysplastic syndromes, which are age-related diseases with assumed mitochondrial defects. Since 2007, her research interest has been on agerelated aspects of viral infections and comorbidities that contribute to severe infections.

Bernadette van den Hoogen studied plant biology at Rijks Hogere Agrarische School (RHAS), Wageningen, The Netherlands. She moved to the field of virology as a research assistant in the Department of Microbiology at the University of Pennsylvania School of Medicine in Philadelphia (under S. R. Ross). She was involved in identifying the receptor for the mouse mammary tumor virus. After accepting a position at Erasmus MC in 1999, she received a Ph.D. in Medicine from Erasmus University in 2004, for her studies on the discovery and characterization of the human metapneumovirus (under R. Fouchier and A. D. Osterhaus). As a senior postdoctoral fellow in the same department, she received a VENI grant from the Dutch Organisation for Scientific Research (NWO) to start her own research line. Her research focuses on the interaction of paramyxoviruses and the innate immune system and the development of therapies against HMPV. She is a frequent reviewer for journals in the field of virology and a member of the editorial board of the Journal of Clinical Virology.

VOL. 24, 2011 Ron Fouchier studied microbiology at the RHAS in Wageningen, The Netherlands, and received a Ph.D. in Medicine from the University of Amsterdam in 1995. He was a postdoctoral fellow at the Howard Hughes Medical Institute, University of Pennsylvania School of Medicine, Philadelphia, PA, from 1995 to 1998, in the laboratory of M. Malim. Achievements of his team include the identification and characterization of several “new” viruses: human metapneumovirus (HMPV), a human coronavirus (hCoV-NL), the severe acute respiratory syndrome coronavirus (SARS-CoV), and a new influenza A virus subtype (H16). He is part of the NIH/NIAID-funded Centers of Excellence in Influenza Research and Pathogenesis and received a prestigious VICI grant from the Dutch Organisation for Scientific Research (NWO) in 2009. He was elected a member of the “Young Academy” of the Royal Dutch Academy of Sciences (2005–2010) and is a member of the Council for Medical Sciences (RMW) and the Central Committee on Animal Experimentation (CCD) to advise the ministry of VWS, Chairman of the Medical and Veterinary Subcommittee of the Committee for Genetic Modification (COGEM), and Vice-Chairman of the COGEM board to advise the ministry of VROM. Dr. Fouchier is active in several WHO and FAO working groups and serves on numerous international advisory boards and on the editorial boards of leading journals. Dr. Fouchier received the Heine-Medin Award in Virology in 2007 and is currently Professor of Molecular Virology at Erasmus MC Rotterdam.

Ralph Tripp is a Professor and holds a Georgia Research Alliance Chair in Vaccine Studies. He was trained in the field of viral immunity under the tutelage of Linda Gooding at Emory University and then with Peter C. Doherty at St. Jude Children’s Research Hospital. Following these postdoctoral programs, he led a research team in vaccine studies for important human viral diseases, particularly RSV, in the Respiratory and Enteric Viruses Branch at the CDC in Atlanta, GA. Professor Tripp now oversees research activities at the Animal Health Research Center (AHRC) at the University of Georgia, which is a biosafety level 2 (BSL2)/BSL3 biocontainment facility where his laboratory develops platform-enabling technologies in pathogen biosensing (using nanotechnology-based approaches), antiviral drugs, and vaccines, using state-of-the-art technologies with an in-house GMP vaccine facility. He is an Editor for Virology and Cellular Immunology and an ad hoc reviewer for several journals. He has received several awards for his work on respiratory viruses.

HUMAN METAPNEUMOVIRUS

753

Rene Alvarez graduated with a degree in biology and received his Ph.D. in microbiology and immunology. During his scientific career, he was a postdoctoral fellow in the Laboratory of the Agricultural Research Service, Athens, GA, and at Southeast Poultry Research and was a research fellow at the Centers for Disease Control and Prevention, National Centers for Infectious Diseases, Division of Respiratory and Enteric Viruses, Atlanta, GA. Between 2004 and 2006, he was Research Assistant Professor at the University of Georgia College of Veterinary Medicine and took various positions up to Associate Director at Alnylam Pharmaceuticals, Cambridge, MA. Currently, he is Supervisory Research Biologist, Principal Investigator, and Head of the Applied Laboratory Science Division, Naval Medical Research Unit, San Antonio, TX. During his career, he has received multiple national scientific awards.

Catherine Manoha-Bourgeois obtained her Ph.D. in Microbiology-Virology in 1991. In 1992, she became Hospital and University Assistant in the Laboratory of Virology, CHU, Dijon, France. Her research activity focused on respiratory syncytial virus. Since 1998, as the Attache´e Scientifique, she has led a small research group that is working on the pathogenesis of respiratory viruses, focusing on metapneumovirus in more recent years. She also directs clinical research on respiratory viruses in the Laboratory of Virology, CHU, Dijon, France.

John Williams received his B.A. with distinction in biology from the University of Virginia and his M.D. from the Medical College of Virginia, where he was elected to the Alpha Omega Alpha medical honor society. He completed a residency in pediatrics at the Children’s Hospital in Pittsburgh and postdoctoral training in pediatric infectious diseases at Vanderbilt University Medical Center. His postdoctoral training was supported by a Pasteur Merieux Connaught award from the Infectious Diseases Society of America. He joined the Department of Pediatrics at Vanderbilt University as an Assistant Professor in 2003. Dr. Williams was awarded the Pediatric Infectious Diseases Society Young Investigator award in 2007. Continued next page

754

SCHILDGEN ET AL.

Oliver Schildgen studied biology at the University of Cologne. His diploma thesis was on the pathogensis of the picornavirus ECHO9. Following his diploma thesis, Oliver Schildgen worked at the University of Cologne’s Institute for Genetics and the Max Planck Institute for Neurological Research in Cologne, Germany, and studied the baculovirus model before performing his Ph.D. thesis work on the woodchuck hepatitis virus in the lab of M. Roggendorf at the University of Essen. Between January 2002 and May 2009, he worked at the University of Bonn with an independent research group that focuses on the epidemiology of new respiratory pathogens. Since June 2009, the group has been based at the Institute for Pathology at the Hospital of the Private University of Witten/Herdecke in CologneMerheim. He has received several national and international awards for his work on newly discovered respiratory viruses.

CLIN. MICROBIOL. REV.

CLINICAL MICROBIOLOGY REVIEWS, Oct. 2011, p. 755–791 0893-8512/11/$12.00 doi:10.1128/CMR.00017-11 Copyright © 2011, American Society for Microbiology. All Rights Reserved.

Vol. 24, No. 4

Serratia Infections: from Military Experiments to Current Practice Steven D. Mahlen* Department of Pathology and Area Laboratory Services, Madigan Healthcare System, Tacoma, Washington INTRODUCTION .......................................................................................................................................................756 HISTORY OF SERRATIA MARCESCENS ..............................................................................................................756 Early History ...........................................................................................................................................................756 Use in Medical Experiments .................................................................................................................................757 Military Use as a Tracer Organism.....................................................................................................................758 NOMENCLATURE AND TAXONOMY OF THE GENUS SERRATIA ..............................................................760 Taxonomy of S. marcescens ....................................................................................................................................760 Taxonomy of Other Serratia Species ....................................................................................................................761 Genomics..................................................................................................................................................................762 NATURAL DISTRIBUTION OF SERRATIA SPECIES ........................................................................................762 HUMAN INFECTIONS CAUSED BY SERRATIA SPECIES ...............................................................................763 S. marcescens ............................................................................................................................................................763 Historical review of infections caused by S. marcescens (1900 to 1960)......................................................764 Opportunistic infections caused by S. marcescens ..........................................................................................766 (i) Opportunistic infections in adult patients ............................................................................................766 (ii) Opportunistic infections among pediatric patients.............................................................................768 Ocular infections caused by S. marcescens ......................................................................................................768 S. liquefaciens ...........................................................................................................................................................769 S. ficaria....................................................................................................................................................................771 S. fonticola ................................................................................................................................................................771 S. grimesii .................................................................................................................................................................772 S. odorifera................................................................................................................................................................772 S. plymuthica ............................................................................................................................................................772 S. quinivorans ...........................................................................................................................................................773 S. rubidaea ................................................................................................................................................................773 VIRULENCE FACTORS OF SERRATIA SPECIES ..............................................................................................773 The S. marcescens RssAB-FlhDC-ShlBA Pathway ..............................................................................................774 Quorum Sensing in Serratia Species ....................................................................................................................774 Enzymes Produced by Serratia Species ................................................................................................................774 ANTIMICROBIAL RESISTANCE OF SERRATIA SPECIES ..............................................................................775 Typical Resistance Patterns of Serratia Isolates.................................................................................................775 Aminoglycoside Resistance in Serratia .................................................................................................................776 ␤-Lactam Resistance in Serratia Species.............................................................................................................776 Chromosomal AmpC ␤-lactamases of Serratia species .................................................................................776 Carbapenem resistance in Serratia species .....................................................................................................777 ESBLs in Serratia species.......................................................................................................................778 Quinolone Resistance in Serratia Species............................................................................................................778 Resistance to the Tetracyclines in Serratia Species ...........................................................................................779 Trimethoprim-Sulfamethoxazole Resistance in Serratia Species .....................................................................780 Treatment of Serratia Species Infections .............................................................................................................780 LABORATORY IDENTIFICATION OF SERRATIA SPECIES ...........................................................................780 Phenotypic Identification .......................................................................................................................................781 Cultural and microscopic characteristics........................................................................................................781 Identification of S. marcescens...........................................................................................................................781 Identification of Serratia species .......................................................................................................................782 (i) S. liquefaciens..............................................................................................................................................782 (ii) S. grimesii...................................................................................................................................................782 (iii) S. proteamaculans.....................................................................................................................................782 (iv) S. quinivorans............................................................................................................................................782 (v) S. ficaria .....................................................................................................................................................782 (vi) S. fonticola .................................................................................................................................................783

* Mailing address: Department of Pathology and Area Laboratory Services, Madigan Healthcare System, 9040 Jackson Ave., Tacoma, WA 98143. Phone: (253) 968-1925. Fax: (253) 968-1068. E-mail: steven [email protected]. 755

756

MAHLEN

CLIN. MICROBIOL. REV.

(vii) S. rubidaea ...............................................................................................................................................783 (viii) S. odorifera..............................................................................................................................................783 (ix) S. plymuthica.............................................................................................................................................783 (x) S. entomophila ............................................................................................................................................783 (xi) S. glossinae ................................................................................................................................................783 (xii) S. nematodiphila ......................................................................................................................................783 (xiii) S. ureilytica .............................................................................................................................................783 Molecular Identification ........................................................................................................................................783 ACKNOWLEDGMENTS ...........................................................................................................................................783 REFERENCES ............................................................................................................................................................783 INTRODUCTION Members of the genus Serratia, particularly the type species Serratia marcescens, cause important infections in humans, animals, and insects. Taxonomically, the genus Serratia is confusing, and currently there are 14 recognized species, with 2 subspecies, in the genus (Table 1). This paper describes the colorful history of S. marcescens and details clinical infections caused by S. marcescens and other members of the genus. First described in 1819, S. marcescens was thought to be a nonpathogen for years, although sporadic reports in the medical literature implicated that the organism could cause opportunistic infections. Since many strains of S. marcescens have red pigment, and the organism was assumed to be nonpathogenic, it was used as a tracer organism in medical experiments and as a biological warfare test agent. In a now-famous expose´, the U.S. government released S. marcescens over both civilian population centers and military training areas from the late 1940s to the mid-1960s in the hopes of gathering data on the potential spread of bioterrorism agents used against the United States. These experiments were unearthed by investigative journalism in the mid-1970s, prompting a congressional investigation that studied U.S. government testing on the public. In the meantime, S. marcescens was revealed to be a pathogen capable of causing a full spectrum of clinical disease, from urinary tract infections (UTIs) to pneumonia. S. marcescens is now an accepted clinical pathogen, and multiantibiotic-resistant isolates are prevalent. Many of the other members of the genus, though, are rarely isolated in clinical microbiology labs and

hence may not be recognized readily by laboratory personnel. The purpose of this review is to give perspective on the history of S. marcescens, provide an update on the taxonomy of the genus Serratia, discuss the natural habitats of the bacteria in this genus, update infections that members of the genus Serratia cause, particularly in humans, and describe the primary identifying characteristics of these organisms. HISTORY OF SERRATIA MARCESCENS Early History In early July 1819, a phenomenon occurred in the province of Padua, Italy, that disturbed many of the peasants in the area, particularly in the town of Legnaro (37, 264). This particular summer had been warmer and more humid than normal, and the polenta, a dish of cornmeal mush made by many families, turned red. Superstitious peasants were fearful of the “bloody polenta,” which was believed to be diabolical in origin. Families refused to stay in homes where the discolored polenta was kept, and one farmer asked for a priest to free his home from “evil spirits” (37, 264). The police were asked to investigate, and they appointed a commission of professors from the University of Padua to assist (37, 264). Bartolomeo Bizio, a pharmacist, studied the phenomenon independently of the University of Padua commission. Bizio conducted experiments wherein he concluded that the red-pigmented polenta was a natural phenomenon in an anonymous paper he authored in August 1819 (37, 49, 264). Bizio successfully cultivated the

TABLE 1. Currently accepted species and subspecies in the genus Serratia Organism

Yr described 关reference(s)兴

Habitat

S. S. S. S. S.

entomophila ficaria fonticola glossinae grimesii

1988 1979 1979 2010 1983

(169) (167) (145) (146) (163)

Insects (Costelytra zealandica) Plants, insects (fig-fig wasp cycle) Water Insects (Glossina palpalis gambiensis) Water, soil

S. S. S. S. S. S. S. S. S. S.

liquefaciens marcescens subsp. marcescens marcescens subsp. sakuensis nematodiphila odorifera plymuthica proteamaculans quinivorans rubidaea ureilytica

1931 1823 1998 2009 1978 1896 1919 1982 1940 2005

(158) (37, 264) (109) (425) (165) (162) (291) (163) (363) (36)

Water, soil, animals, insects, plants Water, soil, animals, insects, plants Water Nematodes (Heterorhabditidoides chongmingensis) Plants Water, animals, insects, plants Water, soil, animals, insects, plants Water, soil, animals, insects, plants Water, plants Water

Pathogenicity

Insects Humans Humans Not reported Not reported (organism has been isolated from human specimens) Humans, insects Humans, animals, insects Not reported Not reported Humans Humans Insects, plants Humans Humans Not reported

VOL. 24, 2011

organism on fresh polenta in these and subsequent experiments and found that reddish discoloration of the polenta could occur in less than 24 h (37, 49, 264). Bizio did not officially publish his results until 1823, when he wrote a letter to Angelino Bellani, a priest, defending his original anonymous article from a paper written by Pietro Melo, Director of the Botanical Garden at Saonara (49). Melo contended, in a paper he wrote in 1819 after he also investigated the phenomenon, that the discolored polenta was due to spontaneous fermentation that turned the polenta into a “colored mucilage” (49, 144). In his 1823 paper, Bizio determined that the cause of the red polenta was an organism he believed to be a fungus that he named Serratia marcescens, after the Italian physicist Serafino Serrati, who pioneered early work on steamboats (37, 49, 264). His description of the genus Serratia was “small, stemless fungi; hemispherical capsules occurring in clusters,” and his description of S. marcescens was “a very thin vesicle filled at first with a pink, then with a red fluid” (37, 49, 144, 264). Bizio observed that small red spots would appear on the cornmeal mush, get larger, and eventually coalesce into a reddish mass of gelatin. These red spots—colonies—apparently looked like “stemless fungi” (49, 144). At the same time that Bizio was conducting his independent investigation, Vincenzo Sette accompanied the University of Padua commission. He came to a similar conclusion as Bizio— that the discolored polenta was a result of a natural process. He presented his data on 28 April 1820 but was not able to publish his findings until 1824. Sette named the causative agent Zaogalactina imetrofa, and he also thought that the organism looked like a fungus (49). Then, in 1848, the naturalist Christian Gottfried Ehrenberg investigated red spots that appeared on a cooked potato in Germany. This discoloration was similar to that seen in the red polenta in Italy; however, Ehrenberg was initially unaware of this. He later read Sette’s published results and concluded that this was probably the same phenomenon. Ehrenberg studied the discolored material under a microscope, and with the improved optics of the time, he saw more detail than the researchers in 1819 were able to see. Ehrenberg noticed actual oval cells in the material, believed that the cells were motile, and stated that they divided longitudinally by fission. In addition, he reported seeing flagella. Because of all of these characteristics, he thought the cells were animals and named the agent Monas prodigiosa (49, 144). Over the course of many years, this organism was described by many different names, and taxonomically it is one of the most complicated organisms that has been described. The now accepted name of S. marcescens was formally adopted in 1980, when the first “Approved Lists of Bacterial Names” was published (358). While Ehrenberg is himself part of the history of the discovery of S. marcescens, he also looked back at the history of the organism and uncovered a much deeper, ancient past (49, 144). Ehrenberg and other investigators described the propensity of S. marcescens to grow on starchy foods, such as bread, and how this growth could be mistaken for fresh blood, especially in times before microorganisms were understood (49, 144). The first reference of “blood” appearing on bread seems to be during the siege of Tyre in 332 B.C. and describes how a seer of the attacking Macedonians said that the “blood” flowing out

SERRATIA INFECTIONS

757

of the bread foretold of the fall of Tyre (49, 144, 176). The Macedonians then went on to take the city (144). Many other events of miraculous, “bleeding” bread have apparently occurred throughout history, many of which are associated with the Host of Christianity. References to the “bleeding” of the Host are not entirely ancient; Breed and Breed described an incident that presumably occurred in Naples in 1910 or 1911 of “bleeding Host” in one of the local churches (49). There are many excellent reviews that cover the ancient history of S. marcescens and “bleeding bread,” including those written by Breed and Breed (49), Gaughran (144), Harrison (176), and Yu (419). Use in Medical Experiments In the late part of the 19th century, William Coley, an oncologist, developed a formula consisting of Streptococcus pyogenes and S. marcescens that he used to treat sarcoma (219). This treatment, called by names such as Coley’s fluid, Coley’s vaccine, Coley’s toxins, and mixed bacterial vaccine, was first used in patients in 1893 by Coley and continued to be used into the 1960s in the United States (219, 413). This preparation was also used in many other countries, and the German pharmaceutical company Su ¨dmedica sold Coley’s toxins under the trade name Vaccineurin until 1990 (219). The efficacy of the treatment has been called into question, but Coley claimed up to 10% cure rates for various types of sarcoma (219). Meanwhile, the first of several medical experiments with S. marcescens as an indicator or tracer organism was conducted by M. H. Gordon in 1906; thus, while the name of the organism was still in question, the pigment characteristics were well known. Gordon was asked to investigate the atmospheric hygiene of the House of Commons in Britain after a recent outbreak of influenza had occurred among the members (9). Gordon, in a now famous experiment, set empty petri plates around him in an empty House of Commons and gargled a liquid culture of S. marcescens to determine the spread of the organisms while delivering passages from Shakespeare (10). S. marcescens colonies were found on plates far enough away from Gordon to show that microorganisms can be spread from speech in addition to being spread by coughing and sneezing (10). Gordon apparently did not become ill from his experiment (10). The next tracing experiment occurred just after World War I. On 15 July 1919, Lieutenant Colonel James G. Cumming and Captain J. W. Cox, both Medical Corps officers of the U.S. Army, sprayed the throats, mouths, and lips of five U.S. Army “donor” soldiers with S. marcescens. The donors were then instructed to eat and then wash their eating utensils and mess kit in warm water. Following this, five unsprayed “recipient” soldiers washed their utensils and mess kits in the same warm water before eating. Transmission of S. marcescens was documented in various percentages from tonsil, tooth, and lip swabs taken from the recipient soldiers, from the mess kits and mess kit wash water, and from the hands of the recipient soldiers. Over the next week, Cumming and Cox conducted two more similar S. marcescens transmission experiments with other donor and recipient soldiers (96). The organism was used as a tracer organism by investigators in medical fields as well. In 1937, Burket and Burn spread S.

758

MAHLEN

marcescens on the gums of dental patients from the outpatient clinic at New Haven Hospital, CT, prior to tooth extraction, in an attempt to show that drawing teeth releases bacteria into the circulatory system. Burket and Burn drew blood cultures from the volunteers after painting their gums and isolated S. marcescens from 7.5% of the cultures (4/53 cultures) in one set of experiments and from 37.8% of the cultures (14/37 cultures) in another set of experiments. The authors concluded that “the use of Serratia marcescens in the present study demonstrated that organisms in the gingival crevice can be forced into the vascular system during extraction” (56). Similar experiments were conducted with S. marcescens in 1949 by McEntegart and Porterfield at the University of Liverpool, with 41.4% (12/29 cultures) recovery of S. marcescens from blood cultures after the organism was applied to the teeth before extraction (260). In an attempt to test equipment designed to remove bacteria from air and to show that S. marcescens could act as a human pathogen, Captain Tom Paine of the U.S. Army conducted an experiment on 2 October 1945, at Camp Detrick, MD, in which he exposed four individuals to about 2,000,000 viable S. marcescens cells per cubic foot of air for 2 1/2 h. Two of the men in the experiment had previously been exposed to S. marcescens by accident in another test of equipment designed to remove bacteria from air. Each of the men was admitted to a hospital and monitored. A few hours after the experiments, each of the subjects developed various signs and symptoms, including body aches, malaise, “smarting of the eyes,” and green sputum production. Three of the subjects had fever and chills, and two of the subjects still had fever at 24 h postexposure. Four days after the experiment, all of the subjects were asymptomatic (292). Paine does not address whether the men in the experiment were military personnel or civilian volunteers. Another set of medical experiments using S. marcescens was conducted at Harvard in 1957 by Kass and Schneiderman. These investigators applied S. marcescens-moistened gauze sponges to different areas of the glans penis of two male patients and to the vulva of a female patient. Each patient had an indwelling catheter, and all were semicomatose. The authors then collected urine from the patients at different times to determine if the presence of the indwelling catheter could facilitate entry of S. marcescens into the bladder. Urine that was collected immediately and 24 and 48 h after application of S. marcescens was sterile; however, S. marcescens was cultured from urine collected at 3 and 4 days postapplication (212). Next, Waisman and Stone wrote a paper in 1958 describing the “red diaper syndrome,” the appearance of S. marcescens in soiled diapers of a female baby born in 1954 at the University of Wisconsin. The parents noticed that soiled diapers that had been rinsed with plain water before being placed in a receptacle provided by a commercial diaper laundry service turned red. This first occurred 3 days after the infant had been discharged from the newborn nursery, and after a week, about one-third of the diapers became red after being placed in the receptacle. At this point, the stool of the infant was cultured and S. marcescens was recovered. Although the baby never had signs or symptoms of illness, physicians treated her with oral sulfasuxidine. Diapers that followed treatment were less red, but the organism persisted in the baby’s intestinal tract for several months. The baby was 2 1/2 years old at the time the paper was written, and no red diapers were observed at that

CLIN. MICROBIOL. REV.

time. The source of this “red diaper syndrome” was initially a mystery. The other parents who had infants born at the same time and who also stayed in the same newborn nursery were contacted, and red diapers were not observed by any of them. It was learned, however, that a biomedical laboratory that was within 500 yards of the hospital had been using S. marcescens in aerosol experiments. Apparently, live organisms were used in the tests and allowed to escape into the air around the laboratory. Another laboratory in an adjoining building reported S. marcescens as an airborne contaminant. The S. marcescens isolate used by the biomedical lab in the aerosol experiments was compared to the patient’s isolate and the contaminant from the other lab, and all three had the same antigenic type (399). Thus, it is more than likely that the baby’s S. marcescens gastrointestinal colonizer came from the strain used in the aerosol experiments. Apparently, the use of S. marcescens as a tracer organism in dental and medical research was common enough that Thayer wrote a paper in 1966 describing the pathogenic potential of the organism, since human infections had started appearing in the literature for several years under the different names of the organism (377); he felt that using the organism as a tracer in human research was open to debate. In 1970, Whalen wrote a short letter stating that laboratory manuals of the time still described procedures for applying S. marcescens to hands and then having students shake hands in an attempt to show how microorganisms can be dispersed (406). By the early 1970s, it was becoming clear that S. marcescens could be a pathogen (1, 16, 34, 101, 139, 144, 172, 177, 294, 302, 314, 324, 407), but for years before that, the organism was thought to be a nonpathogen and an ideal tracer organism. In fact, events in the 1970s eventually detailed just how often S. marcescens was used as a tracer organism, and not in just medical experiments. Military Use as a Tracer Organism In 1977, the U.S. Senate Subcommittee on Health and Scientific Research held hearings that described biological warfare tracer organism tests that the U.S. military had conducted on military bases and the general population from the 1940s through the 1960s (11). One of the organisms used in the tests was S. marcescens. Except for Cumming and Cox studying transmission of S. marcescens among soldiers after World War I (96), it is not precisely known when this organism was first used by militaries in tracing experiments. The earliest reference appears in the 1930s, as described by Henry Wickham Steed. Steed, a respected British journalist and previous editor of The Times, wrote an article published in 1934 in the periodical The Nineteenth Century and After in which he alleged that Germany was actively involved in biological warfare experimentation (191, 322). Steed described documents that he received from sources that contained notes with experiments conducted by the Germans in Paris first on 18 August 1933 (191). According to Steed, German agents released S. marcescens aerosols near ventilation shafts at various locations of the Paris Me´tro, including the Place de la Concorde and probably other sites (191). Other agents had placed plates on various Me´tro platforms, Steed maintained, such as at the Place de la Re´publique, and counted the number of S. marcescens colonies in an

VOL. 24, 2011

attempt to determine the efficiency of aerosol dispersal of a potential biological warfare agent. The documents that Steed obtained also allegedly describe aerial release experiments of S. marcescens at the Berlin Templhof airfield while the Paris Me´tro releases were occurring. In addition, other experiments were alleged to have taken place at other locations in France. The documents that Steed received apparently no longer exist, and it is probable that he destroyed them in 1939. The authenticity of the notes, including the obtained microbiological data, has been called into question. For example, data were collected from the Pasteur Station, with the note “95,778 colonies were counted!!! The result was checked an hour later and 91,389 colonies were counted”; thus, the notes seem to indicate that colonies were counted an hour after release in at least one case, and the colony counts are probably too precise as well. Some consider that the documents that Steed received were forgeries. The French took Steed’s article very seriously at the time; Germany denied the report (191). Then, in the mid-1970s, came the news that triggered the U.S. Senate Hearings before the Subcommittee on Health and Scientific Research of the Committee on Human Resources (11). The Long Island newspaper Newsday published a report in their 21 November 1976 paper that described tests that the U.S. government had conducted on the population of San Francisco in 1950 and also in the New York City subway system to determine how vulnerable these cities were to a biological warfare weapon attack and also to determine the viability of organisms used in these tests (11, 84). The report said that one person had died of an infection caused by the same organism, S. marcescens, used in the tests, and that at least five other patients had been ill with S. marcescens infections (11, 85). A month later, on 22 December 1976, the Washington Post reported several other instances of tests involving S. marcescens and other microbes (Bacillus globigii and Aspergillus fumigatus) at both military installations and U.S. cities (11). The U.S. Army did acknowledge that testing had been conducted with S. marcescens at eight locations on 15 December 1976, so the article in the Washington Post probably utilized that information (11). On 23 December 1976, the Atlanta Constitution reported eight locations where tests were run, with dates: the Pentagon, Washington, DC (1950); San Francisco (1950); Mechanicsburg, PA (1951); Key West, FL (1952); Fort McClellan, AL (1952); Panama City, FL (1953); Point Mugu-Port Hueneme, CA (1956); and New York City (1966) (11). The hearings, which took place on 8 March 1977 and 23 May 1977, revealed that S. marcescens had been tested at “public domain” sites, i.e., civilian population areas, a total of at least 7 times, from 18 August 1949, in Washington, DC, until March 1968, in Hawaii. In addition, S. marcescens was tested against non-public-domain sites (government and military facilities) at least 29 times, ranging from December 1950, at Naval Amphibious Base, Little Creek, VA, to 16 October 1968, at Edwards Air Force Base, CA (11). Other agents, such as B. globigii and A. fumigatus, were used in many of the same tests and also in other tests on other sites (11). In 1981, the grandson of the individual who died from the S. marcescens infection in San Francisco in 1950 (11, 407) sued the U.S. government over the testing that allegedly introduced S. marcescens to his grandfather (85). The tests in September 1950 were accomplished by U.S. Navy ships releasing aerosols

SERRATIA INFECTIONS

759

containing S. marcescens and B. globigii off the coast of San Francisco; winds then carried the organisms inland (85, 419). Collection stations were established at several inland positions, and B. globigii was readily isolated, probably because it is a spore former (85). However, S. marcescens was not isolated as readily; it was thought that perhaps the organism had lost its pigmentation and that that was why it was not found by the collection stations (85). In 1951, Wheat and others wrote a paper describing a cluster of probable S. marcescens urinary tract infections that occurred in patients at Stanford (407). The authors stated that up until this point, they had not isolated S. marcescens at their institution, and that one of the patients had died as a result of an S. marcescens infection. About a month before the tests were conducted, the patient who eventually died from the S. marcescens infection had developed acute urinary retention while dealing with arteriosclerotic heart disease. A catheter was placed, and a month later his prostate was surgically removed. The patient soon developed a urinary tract infection, and a red-pigmented Gram-negative rod was isolated from his urine. He was eventually admitted, and the same isolate was recovered from blood cultures. The patient died on hospital day 21, of endocarditis. Around the September-October 1950 time frame, four other red-pigmented Gram-negative isolates were recovered from different patient urine cultures, and then six more were recovered from November 1950 to February 1951 (407). The strains isolated at Stanford were not archived and were never compared to the strain used in the testing. Each patient in the Stanford cluster had urinary tract complications, and it is not unusual to see S. marcescens nosocomial outbreaks among similar populations (407, 419). Since proof could not be established that the same strain of S. marcescens that caused the death of the patient in San Francisco in 1950 was the strain used in the vulnerability tests, the judge did not rule in favor of the patient’s family (86). Farmer and others, in response to concerns in the U.S. press and a paper by Severn that discussed that a potential reason for more reported cases of S. marcescens infections in the United States than in other countries was the U.S. government experimentation, conducted a thorough investigation of S. marcescens strains that had been collected at the Centers for Disease Control and Prevention (CDC) from 1950 through the publication date of their paper in 1977 (129, 345). The CDC obtained the S. marcescens isolate that was used in vulnerability testing from Fort Detrick, where the isolate had been prepared for use in tests, in 1977 and found that it had the same type (S. marcescens 8 UK, biotype A6, serotype O8:H3, phage type 678) characteristics as isolates that they had preserved from 1957 and 1969 (129). Thus, the isolate used in the population vulnerability tests was stable (129). Over 2,000 S. marcescens cultures were biotyped in the study, and only 20 were of biotype A6, which is a rare biotype (129, 162). There were 7 U.S. isolates that were biotype A6, but only one that was serotype O8:H3; Farmer and others do not relate whether this was a clinical isolate or not, although biotype A6 is usually isolated from the environment (129, 159). In addition, the CDC serotyped over 3,000 S. marcescens isolates during the period of this study and found only 7 O8:H3 serotypes; it is not mentioned if any of these were isolated from clinical specimens (129). By 1977, there were more than 100 outbreaks of S. marcescens in the United States, and none had the same strain

760

CLIN. MICROBIOL. REV.

MAHLEN

characteristics as the isolate used in the vulnerability tests (129). Thus, the authors concluded that the strain used in testing was not an important cause of morbidity and mortality in the United States (129). Several sources make for interesting reading. The Hearings before the Subcommittee on Health and Scientific Research of the Committee on Human Resources that describe the congressional investigation are publically available (11). Leonard A. Cole’s book Clouds of Secrecy discusses the San Francisco S. marcescens release, the trial involving the grandson of the individual who died of the S. marcescens endocarditis described by Wheat and others, and other events concerning government-sanctioned testing over public areas (84). Yu’s 1979 review paper also provides a detailed summary of the military use of S. marcescens as a dispersal agent (419). NOMENCLATURE AND TAXONOMY OF THE GENUS SERRATIA Taxonomy of S. marcescens S. marcescens has one of the most confusing taxonomies in the bacterial world, and part of the confusion no doubt stems from the uncertainty about whether the early descriptions of the organism by Bizio, Sette, Ehrenberg, and others were redor pink-pigmented yeast or bacteria; microorganisms such as Rhodotorula spp., Methylobacterium spp., Roseomonas spp., Azospirillum spp., and others could all potentially have been thought to be the same organism since the 19th century. Also, other red-pigmented Serratia species, such as S. rubidaea and S. plymuthica, could have been confused in some cases with S. marcescens, especially since most members of the genus are found in the environment (Fig. 1 shows typical red pigmentation of S. marcescens on different types of agar media). In 1920, Winslow and others published the Final Report of the Committee of the Society of American Bacteriologists on Characterization and Classification of Bacterial Types, and they named the organism Erythrobacillus prodigiosus, following a report by Louis Fortineau in 1904 (411). This was challenged initially in the 1st edition of Bergey’s Manual of Determinative Bacteriology, in 1923, when Breed wrote that the name S. marcescens took precedence over all other proposed names (49). Breed and Breed had performed an extensive study of the history of S. marcescens and uncovered Bizio’s early work (49). Up until the time that Breed used the name S. marcescens in the 1st edition of Bergey’s Manual, there had been 17 other names used for the organism (144). After Bergey’s Manual of Determinative Bacteriology was first published, three more names were used for S. marcescens: Salmonella marcescens, Salmonella prodigiosum, and Chromobacter prodigiosum (144). C. prodigiosum, in particular, was used commonly until the 1950s. Cowan maintained in 1956 that Bizio had studied a yeast and that the resolving power of the microscopes available at the time was not adequate to see a typical Gram-negative bacillus but was probably adequate to see yeast cells (92). Thus, Cowan felt that S. marcescens should not be the official name (92). Despite Cowan’s objections, the International Code of Nomenclature of Bacteria and Viruses, Bacteriological Code (1958) published S. marcescens as the official name of the organism (144). Several years later, Gaughran wrote in Bizio’s defense that van Leeu-

FIG. 1. Red-pigmented colonies of S. marcescens on MacConkey agar (A), tryptic soy agar (B), and tryptic soy agar with 5% sheep blood (C). The cultures were incubated at 35°C for 18 h. The MacConkey agar plate was incubated in ambient air, and the other two plates were incubated in 5% CO2. Each plate was inoculated with the same strain of S. marcescens, which was isolated from a case of endophthalmitis.

VOL. 24, 2011

SERRATIA INFECTIONS

wenhoek saw individual bacteria in 1683 with his antiquated microscope, so it was certainly possible for Bizio to see a bacterium such as S. marcescens in 1819 with the improved optics of the time (144). Gaughran also concluded that Bizio’s description of the colonies seems more likely to fit the description for bacterial colonies than yeast cells (144). Every edition of Bergey’s Manual of Determinative Bacteriology used the name S. marcescens throughout the 1900s, and S. marcescens was established as the official name in 1980, when “Approved Lists of Bacterial Names” was published under the direction of the International Committee for Systematic Bacteriology (358). Publication of the approved lists of bacterial names also established 1 January 1980 as the new date for determining priorities for names of new taxa, replacing the previously used date of 1 May 1753 (358). In particular, the reviews by Breed and Breed (49) and Gaughran (144) provide comprehensive summaries of the taxonomy of S. marcescens. In 1998, a red-pigmented endospore-forming organism was recovered from a wastewater treatment tank in Saku, Japan (109). At the time, it was reported as a probable Bacillus species, but the DNA G⫹C content resembled that of the genus Serratia (2). Numerous studies by Ajithkumar and others were undertaken to determine the identity of the isolate. The DNA G⫹C content matched that of S. marcescens (58 mol%), and the 16S rRNA gene sequence was 99.6% similar to that of S. marcescens. Transmission electron microscopy was performed on the isolate, and it had endospores and a Gramnegative type of cell structure. The organism produced prodigiosin, the compound responsible for red pigmentation in many strains of S. marcescens, S. plymuthica, and S. rubidaea, and had the same biochemical pattern as S. marcescens (2). The formation of endospores had never before been reported for members of the Enterobacteriaceae, and confirmation of the existence of the endospores is now in question; a member of the Subcommittee on the Taxonomy of Enterobacteriaceae for the International Committee on Systematics of Prokaryotes has so far not been able to identify spores in the isolate (185). Ajithkumar and others, in the paper where they described this endospore-forming isolate of S. marcescens, suggested that the organism may have undergone gene transfer with Bacillus species present in the wastewater in order to acquire the ability to form endospores (2). If gene transfer can occur between S. marcescens and Bacillus species in nature, then perhaps S. marcescens may also readily lose the acquired genes. At any rate, the isolate is considered to belong to a subspecies of S. marcescens, and at this point it is officially known as S. marcescens subsp. sakuensis, while the type strain of S. marcescens is referred to as S. marcescens subsp. marcescens (2; http://www .bacterio.cict.fr/s/serratia.html).

Taxonomy of Other Serratia Species Confusion exists about the nomenclature of other Serratia species as well; see Table 1 for dates that Serratia species were described. S. liquefaciens, S. proteamaculans, S. quinivorans, and S. grimesii belong to the S. liquefaciens complex (159). S. liquefaciens was first described in 1931 by Grimes and Hennerty, as Aerobacter liquefaciens (158). In 1963, this organism was placed in the genus Enterobacter (125). Since this

761

organism was phenotypically similar to S. marcescens, E. liquefaciens was reassigned as S. liquefaciens in 1973 (126). S. proteamaculans was first identified in 1919, when Paine and Stansfield recovered it from cases of leafspot disease on the tropical flowering plant Protea cynaroides (291). At the time, they named it Pseudomonas proteamaculans, and the organism has since been renamed several times, including both Bacterium proteamaculans and Phytomonas proteamaculans in 1930 (166). By 1948, Burkholder had renamed the organism Xanthomonas proteamaculans (57), and then Dye classified it as Erwinia proteamaculans in 1966 (118). This name held until 1974, when Lelliott wrote that the organism was possibly an Enterobacter species but should be excluded from the genus Erwinia because of some of its biochemical characteristics (236). Then, in 1978, Grimont and others studied Erwinia proteamaculans and concluded that it was synonymous with a strain of Serratia liquefaciens (166). The “Approved Lists of Bacterial Names” in 1980 listed both Serratia proteamaculans and S. liquefaciens as separate species (358), and in 1981 Grimont and others provided evidence that both were indeed distinct (168). In 1982, Grimont and others determined that a biogroup of S. proteamaculans should be designated a subspecies, S. proteamaculans subsp. quinovora (163). Most recently, Ashelford and others proposed in 2002 that this subspecies be elevated to a distinct species, Serratia quinivorans (20). In 1983, Grimont and others described S. grimesii after they studied 11 Serratia strains that were isolated from water, soil, and human samples; they named the organism after the Irish bacteriologist Michael Grimes, who first described this group (158, 163). S. rubidaea was originally described by Stapp in 1940 as Bacterium rubidaeum and reassigned as a Serratia species in 1973 (126, 363). It is a red-pigmented organism, and the species epithet is a contraction of the scientific name for the raspberry plant, Rubus idaeus. In 1944, Zobell and Upham described S. marinorubra, a red-pigmented organism they isolated from marine water (427). In 1980, the “Approved Lists of Bacterial Names” determined that both species had the same type strain and thus were homotypic synonyms (358). Since they are homotypic synonyms, the name S. rubidaea has priority (160). Apart from S. marcescens, the oldest member of the genus Serratia is S. plymuthica. It was first identified by Fischer in 1887 as a red-pigmented organism isolated from the water supply of Plymouth, England. It was originally called Bacillus plymouthensis by Dyar in 1895, but he did not validly publish it, so the first published name of this organism was Bacterium plymuthicum, by Lehmann and Neumann in 1896. This organism was then transferred to the genus Serratia in 1948 in Bergey’s Manual and was renamed S. plymuthica (162). The taxonomy of the other currently recognized Serratia species is clearer. S. odorifera was named in 1978 by Grimont and others, who studied 25 similar strains that were isolated mostly from various human specimens (165). S. odorifera is not pigmented and was named for its characteristic potato-like odor (165). Then, Gavini and others found that 20 organisms that had similar characteristics and that were isolated from water were a new species, and they named it S. fonticola in 1979 (145). S. fonticola does not share many of the key characteristics of other Serratia species, such as gelatin hydrolysis

762

CLIN. MICROBIOL. REV.

MAHLEN

different types of aphids and apparently has only recently evolved as a symbiont (55). Patrick Grimont and Francine Grimont have written many papers describing the members of the genus Serratia, including several excellent taxonomy reviews (159–162). Genomics

FIG. 2. Dendrogram of the genus Serratia, constructed using the neighbor-joining method in MicroSeq software. 16S rRNA gene sequences of type strains of the species, as listed in the List of Prokaryotic Names with Standing in Nomenclature at http://www.bacterio.cict .fr/s/serratia.html, were obtained from GenBank. GenBank accession numbers are listed after the species in the dendrogram. The sequence of Proteus mirabilis, used as the outgroup, was from the MicroSeq database. The length of the bar at the top represents a 4.611% difference in 16S rRNA gene sequence.

and DNase production, and it has a lower mol% G⫹C (49 to 52% for S. fonticola, compared to 52 to 60% for other members of the genus Serratia) (159). Because of this, S. fonticola is sometimes thought of as temporarily assigned to the genus Serratia (128), but it is still officially listed as a Serratia species (159; http://www.bacterio.cict.fr/s/serratia.html). By 16S rRNA gene sequence analysis, S. fonticola belongs in the genus Serratia (Fig. 2) (159). S. ficaria was also described in 1979, when 14 related strains that were recovered from figs, caprifigs, fig wasps, and a black ant were studied (167). Next, a Serratia species that caused amber disease in rot grubs was identified and called S. entomophila in 1988 (169). In 2005, S. ureilytica was isolated from river water in West Bengal, India (36). In 2009, a red-pigmented organism was isolated from the intestine of Heterorhabditidoides chongmingensis, an entomopathogenic nematode (425). This isolate, named S. nematodiphila, was also fluorescent (425). The most recent accepted species is S. glossinae, described in 2010 after it was isolated from the midgut of the tsetse fly (Glossina palpalis gambiensis) (146). Most recently, the species “S. symbiotica” was proposed based on DNA and protein phylogenetic studies (54, 230, 330). This bacterium is a secondary symbiont associated with several

To date, only one complete genome has been sequenced for the genus Serratia, that of S. proteamaculans strain 568 (GenBank accession number CP000826). The genome is 5.45 Mbp, with 4,891 genes encoding proteins, and the strain also has one 46-kb plasmid that was sequenced (GenBank accession number CP000827). The genome was sequenced by the U.S. DOE Joint Genome Institute, and the project can be viewed at http://www .ncbi.nlm.nih.gov/sites/entrez?Db⫽genomeprj&Cmd⫽Search &Term⫽txid399741[orgn]. There are several genomes that are in the process of being sequenced. Two different strains of S. marcescens, ATCC 13880 and Db11, are currently being sequenced, by the University of Wisconsin-Genome Evolution Laboratory and the Sanger Institute, respectively. Likewise, two different strains of S. odorifera, 4Rx13 and DSM 4582, have been sequenced and are being assembled. The genome of S. odorifera strain 4Rx13 is 5.36 Mbp, and that of strain DSM 4582 is 5.13 Mbp. Two different strains of “S. symbiotica” are also being sequenced. “S. symbiotica” strain Tucson was sequenced by the University of Arizona and is being assembled. Like those of other symbiotic bacteria, the 2.57-Mbp genome is smaller than that of free-living bacteria. The genome has undergone genetic decay since becoming a symbiote compared to other members of the genus Serratia (55). Another strain, Cinara cedri, is currently being sequenced by Valencia University. Lastly, there are several Serratia strains that are being sequenced that have not yet been named. These strains have been identified from environmental sources or, in a few cases, from human specimens. A complete listing of complete bacterial genome sequences and genomes that are in the progress of being sequenced can be viewed at http://www.ncbi.nlm.nih.gov /genomes/lproks.cgi. NATURAL DISTRIBUTION OF SERRATIA SPECIES Since the appearance of the discolored polenta that Bizio and Sette studied, the red-colored potato that Ehrenberg studied, and the earlier findings of “bloody” bread and Host through the ages, it was apparent that S. marcescens was readily found in the environment. Because of the confusing taxonomic status of the members of the genus, it was not always readily apparent which natural environments the different species were found in. It is known now, however, that Serratia species are commonly found in water and soil and are also associated with plants, insects, and animals. Common habitats of Serratia species are listed in Table 1. Water appears to be a natural environment for several species, including S. marcescens, S. fonticola, S. grimesii, S. liquefaciens, S. plymuthica, S. rubidaea, and S. ureilytica (23, 36, 145, 159–162, 209, 416). S. marcescens, S. liquefaciens, S. proteamaculans, S. grimesii, and S. plymuthica were found in river water in one study, with the predominant species being S.

VOL. 24, 2011

marcescens, followed by S. liquefaciens (160). S. marcescens subsp. sakuensis was originally isolated from the suspended water of a wastewater treatment tank in Japan (2). Many Serratia species are also associated with soil, including S. marcescens, S. grimesii, S. liquefaciens, and S. quinivorans (20, 159, 161). Klein isolated what was probably S. marcescens from cooked meat and fish in the late 1800s from a wholesale mercantile house in London (220). He theorized that the organism contaminated the food products after soil and graves in an adjoining churchyard had been disturbed; the wind had been blowing toward the mercantile house while the work commenced (220). S. marcescens is found naturally in different soil types (23, 142, 161). Perhaps because Serratia species are found in soil, several are associated with plants (161). S. marcescens and S. liquefaciens appear to be the most commonly plant-linked Serratia species and have been isolated from many different types of plants, including grass, tomatoes, green onions, and other vegetables (161). S. quinivorans was isolated from soils associated with plants such as sugar beets (20). It is possible that in some cases soil is the source of organisms such as S. marcescens isolated from plants. In some cases, though, Serratia species are found closely associated with plants and may be important for plant health. For example, S. plymuthica is able to stimulate the growth of plants and suppress soilborne plant pathogens (279). Also, S. liquefaciens, S. plymuthica, and S. rubidaea were associated with the rhizosphere of oilseed rape, and all three demonstrated antifungal properties (208). In addition, S. rubidaea was found associated with marine alga in one study (209). S. proteamaculans was originally identified as a cause of leafspot disease of Protea cynaroides, the king protea, the national flower of South Africa (291). This organism may be the only Serratia species identified that is a phytopathogen, although S. marcescens was reported to cause a hypersensitivity reaction when applied to tobacco and bean leaves (229). One particularly close association of Serratia species with plants is that of S. ficaria and fig trees. S. ficaria has been found in figs in many places in the world, including France, Greece, Sicily, Tunisia, and California (160). S. ficaria has also been recovered from fig wasps, which pollinate Smyrna and Calimyrna figs (160). S. ficaria was recovered from a patient with endophthalmitis in South Australia; the patient kept figs on his property, so it can be assumed that the organism can be recovered from figs in that part of the world as well (25). In addition to an association with plants, Serratia species are also found in insects, and some species are pathogenic to insects. S. entomophila was first found as a cause of amber disease in grass grubs (169), and S. proteamaculans has also been found as a cause of amber disease (151, 170). S. marcescens is pathogenic to at least 70 species of insects (164). S. marcescens, S. plymuthica, S. ficaria, and S. liquefaciens have all been isolated as part of the natural floras of many different kinds of insects, including flies, wasps, termites, and grasshoppers (161). Some of these organisms may also be pathogenic for the same insect varieties (161). Serratia species are also associated with animals and cause important animal diseases. S. marcescens was described in 1958 as a cause of illness in animals, when part of a dairy herd was diagnosed with mastitis (27). There are many other reports of colonization or disease by Serratia species in animals, including

SERRATIA INFECTIONS

763

but not limited to reptiles, rodents, birds, chicks, goats, pigs, fish, and horses (29, 160). Most recently, S. marcescens was identified as the causative agent of white pox disease, a serious threat to the Caribbean elkhorn coral, Acropora palmata (301). It is probable that the S. marcescens strain responsible for white pox disease in A. palmata, which is classified as threatened by the U.S. Endangered Species Act, is of human fecal origin (373). The same S. marcescens strain was isolated from two other coral species and from a marine snail from the same region (373). There are many excellent reviews that cover the natural distribution of Serratia species, including those written by Patrick Grimont and Francine Grimont (159–161). HUMAN INFECTIONS CAUSED BY SERRATIA SPECIES Human infections by members of the genus Serratia, particularly S. marcescens, were not well recognized until the latter half of the 20th century. This is probably due to the challenge of taxonomically describing the genus and to the fact that several species were not identified until the 1970s and 1980s. S. marcescens is now recognized as an important human pathogen; however, many other members of the genus occasionally cause human infections. At this time, S. entomophila, S. glossinae, S. proteamaculans, S. nematodiphila, and S. ureilytica have not been implicated in human infections. In large surveys, Serratia species account for a relatively low percentage of isolates from different types of infections; while it can be assumed that most of these Serratia infections are due to S. marcescens, in some cases the species is not established. In a survey of ICU-acquired infections in European countries by the European Centre for Disease Prevention and Control in 2008, Serratia species represented 2.0% of all bloodstream infections, ranking organisms from this genus as the 10th most commonly recovered organisms from ICU-acquired bloodstream infections (12). A survey from 1997 data on SENTRY Antimicrobial Surveillance Program isolates from the United States, Canada, and Latin America showed that Serratia species were the 12th most common organisms associated with bloodstream infections, accounting for 1.4% of all isolates (107). For ICU-acquired pneumonia cases from Europe in 2008, Serratia species represented 2.8% of all such infections and were the 11th most commonly isolated organisms (12). Data from the SENTRY Antimicrobial Surveillance Program from 2004 to 2008 revealed that Serratia species were isolated from 3.5% of all patients hospitalized with pneumonia. In this survey, the incidence of Serratia from patients with pneumonia in the United States was 4.1%, while the incidence was 3.2% in Europe and 2.4% in Latin America. Overall, Serratia species were the seventh most common cause of pneumonia in hospitalized patients in this study (205). S. marcescens S. marcescens is the most commonly isolated Serratia species in human infections (160, 233). Like many of the other members of the Enterobacteriaceae, S. marcescens has been recovered from a large variety of clinical specimens. S. marcescens causes central nervous system diseases such as meningitis (16,

764

MAHLEN

314), urinary tract infections (140, 231, 376, 407), pneumonia and other respiratory diseases (1, 34, 172, 413), bloodstream infections, including endocarditis (177, 302, 407), and many different types of wound infections (140, 314). In 1979, Yu reviewed a few cases of septic arthritis, osteomyelitis, and endocarditis caused by S. marcescens in heroin addicts (419), typified by a report by Mills and Drew of 19 endocarditis cases in addicts from San Francisco from 1963 to 1974 (268). At the time, it was thought that S. marcescens may be a serious pathogen in drug abusers, but there have not been many reports of S. marcescens infections in this patient population since the 1970s, so these types of infections may be sporadic. At my facility, Madigan Army Medical Center, a U.S. Army health care system that serves active duty military personnel and their dependents, as well as military retirees and their dependents, S. marcescens is the ninth most commonly isolated Gram-negative rod and comprised 214 isolates from 156 different patients from 2005 through 2010 (unpublished data). The most frequent source of isolation was respiratory tract specimens (72 isolates; 33.6%), followed by urine specimens (51 isolates; 23.8%) and various wound culture isolates (49 isolates; 22.8%). During the same period, only one other Serratia species, S. liquefaciens, was isolated from a human specimen at my facility (unpublished data). My hospital is in Pierce County, WA, and in 2009 S. marcescens was the eighth most commonly reported Gram-negative rod from Pierce County hospitals (unpublished data). A large, nationwide survey from Poland from November 2003 to January 2004 revealed that S. marcescens was the fifth most commonly recovered organism of the Enterobacteriaceae family, representing 4% of all Enterobacteriaceae clinical isolates (122). A nationwide survey from Japan from January 2008 to June 2008 showed that S. marcescens caused 6.4% of urinary tract infections; S. marcescens was the fifth most common cause of urinary tract infections in that study (194). In the literature, there has been a very large number of reported hospital-related S. marcescens outbreaks since the 1950s (⬎200). Because there are so many described hospitalassociated outbreaks, it is often assumed that infections caused by S. marcescens are primarily nosocomial in origin. Recently, however, Laupland and others conducted an extensive survey of Serratia infections in Canada and found that 65% of all infections caused by Serratia species were community based. In this report, S. marcescens was the most commonly isolated species, accounting for 92% of all isolated Serratia species (233). The literature, however, is dominated by outbreaks and opportunistic infections caused by S. marcescens. In addition, S. marcescens is an ocular pathogen of note, and not always in hospitalized or immunocompromised patients. Historical review of infections caused by S. marcescens (1900 to 1960). Because of the taxonomic confusion that has existed over the years for members of the genus Serratia, and because S. marcescens is not always pigmented, reviewing early literature for references of S. marcescens infections in humans is somewhat challenging. Most of the papers that describe probable S. marcescens infections of humans from the first 60 years of the 20th century attribute the infections to Chromobacterium prodigiosum, and in some cases, the authors themselves have questioned the identity of the recovered red-pigmented

CLIN. MICROBIOL. REV.

organism (172, 302). Part of this confusion can be attributed to early descriptions of the so-called “chromobacteria group.” The chromobacteria were classified as three different bacteria based on their ability to form pigment; thus, “Chromobacterium prodigiosum” produced pink or red colonies, Chromobacterium violaceum produced a violet pigment, and “Chromobacterium aquatilis” produced yellow or orange colonies (407). In addition, biochemical identification of bacteria at the time was not as sophisticated as modern methods, and molecular methods to resolve discrepancies were not available. Thus, the identity of the causative agent in some of the earlier references to S. marcescens human infections can be questioned. However, these early cases are informative when viewed together and show a framework of the pathogenic potential of this organism, especially with regard to the ability to cause nosocomial infections or infections in immunocompromised patients. Table 2 summarizes reported, probable S. marcescens cases from 1900 to 1960. The first probable case of reported incidence of human infection by S. marcescens was the isolation of a red-pigmented organism, called Bacterium prodigiosum, from the sputum of a patient with a chronic cough, published in 1913 by Woodward and Clarke. The patient was not immunocompromised and was apparently healthy prior to infection but had a persistent cough for 3 years. The patient had noticed that his sputum was red and smelled bad, so he consulted a physician because he feared tuberculosis. The investigators noticed that the pigment of the organism was lessened on subculture (413). This case perhaps represented colonization of the respiratory tract by S. marcescens, not true infection. Another case of S. marcescens isolated from the sputum of a patient with pneumonia was described in the French literature in 1936 (1). The next published case in the English literature of S. marcescens infection in a human was a case of meningitis in a U.S. Army soldier in 1942. The soldier had previously been diagnosed with syphilis, and in July 1941, he had a diagnostic lumbar puncture performed. Antisyphilitic treatment was continued, and the soldier had another lumbar puncture procedure in February 1942. The soldier complained of having coldlike symptoms, including a cough, at this time. In 3 days, the soldier had signs and symptoms of meningitis, and red-pigmented, motile, Gram-negative bacteria that were thought to be S. marcescens were isolated from cerebrospinal fluid (CSF) from repeated lumbar punctures. The patient improved and was discharged in May 1942. The source of S. marcescens in this case is unclear, but it may have been introduced nosocomially when the patient underwent one of the diagnostic lumbar puncture procedures (16). Wheat and others described several nosocomial UTIs, with a case of fatal endocarditis, caused by S. marcescens in San Francisco in 1951. A year before, the first probable case of S. marcescens UTI was described by Gurevitch and Weber, who described a 61-year-old male who was admitted in December 1948 in Jerusalem, Israel, with acute bronchopneumonia. A week after admission, the patient had dysuria, and a red-pigmented organism, identified as “Serratia,” was recovered from the urine along with Escherichia coli and Staphylococcus aureus. Pure cultures of Serratia were isolated four more times from the patient’s urine over the next 15 days. The authors found that the isolate was similar to S. marcescens but had some differences. For exam-

VOL. 24, 2011

SERRATIA INFECTIONS

TABLE 2. Summary of S. marcescens infections from 1900 to 1960a Yr of report

1913 1936 1942

1948 1950–1951

1951

1951–1952

1953 1953

1957 1957 1960

Comments

Reference

Previously healthy patient with chronic cough; red-colored sputum; redpigmented organism recovered In the French literature; recovered from sputum of patient with pneumonia Meningitis from a U.S. Army soldier who had previously had a diagnostic lumbar puncture performed; redpigmented organism recovered UTI in patient admitted with acute bronchopneumonia; red-pigmented organism recovered Outbreak of 11 cases of UTI; 1 patient died from endocarditis, presumably from the same isolate; all strains were red pigmented Fatal sepsis in patient who had a gastrectomy because of a duodenal ulcer; red pigmented bacterium recovered Outbreak of 12 cases in a pediatric ward in Israel; several types of infections, including wound infections, skin lesions, meningitis, otitis, and shoulder joint arthritis; 1 fatal case of meningitis in a neonate; outbreak traced to bottle of 5% glucose in saline; all isolates were red pigmented Fatal endocarditis in a patient from the former Gold Coast (Ghana); red-pigmented organism recovered Patient had red-colored sputum after coughing, simulating hemoptysis; red-pigmented organism recovered; similar to 1913 Woodward and Clarke case Empyema in patient with right spontaneous pneumothorax; redpigmented organism recovered Pseudohemoptysis; red-pigmented organism recovered Pneumonia in patient with tuboovarian abscess; red-colored sputum; red-pigmented organism recovered

413 1 16

172 407

302

314

177 139

294 324 34

a Infections were assumed to be caused by S. marcescens based on the recovery of red-pigmented organisms.

ple, they stated that their isolate grew at 37°C but that S. marcescens does not; it is now known that S. marcescens will certainly grow at 37°C. Gurevitch and Weber named their isolate “Serratia urinae,” but it certainly could have been S. marcescens (172). The source of the organism in this case was not clear, but it seems to be nosocomial in origin. In 1952, a case of S. marcescens fatal sepsis was reported by Patterson and others for a 63-year-old male patient with a history of a gastrectomy because of a duodenal ulcer. The previous year, the patient was admitted with hematemesis, melena, and weakness; by hospital day 29, the patient became septic and S. marcescens was recovered from several blood cultures. The patient was treated at different times with aureomycin, chloramphenicol, and streptomycin and eventually died

765

on hospital day 51, despite therapy. The authors stated that the pink-to-red-pigmented isolate resembled the descriptions of both “Chromobacterium prodigiosum” and S. plymuthicum, but they used the recommended taxonomy of the time to name the organism. Interestingly, Patterson and others reported that UTIs were the most common clinical manifestation of S. marcescens in humans. They did not cite a specific reference but cited unpublished data from J. Draper from Bellevue Hospital, NY, who found 2 cases of UTI caused by “chromobacteria” out of 100 UTI cases (302). No data are presented as to the actual identity of the chromobacteria that caused these UTI cases. Also in 1952, Rabinowitz and Schiffrin reported a fatal case of S. marcescens meningitis in a 4-month-old child in Israel. The infant had been admitted originally for enteritis in late 1951 and was initially treated with penicillin and sulfaguanidine. Three days later, the infant developed meningitis and S. marcescens was recovered from CSF. Therapy had been switched to streptomycin after Gram-negative rods were observed in the CSF, but the infant died. This case occurred among a series of S. marcescens infections from the same pediatric ward at the same hospital in Jerusalem. Previously, S. marcescens was isolated from wound infections from two other children. After the meningitis case, nine other S. marcescens infections occurred in children from the same ward between December 1951 and January 1952; infections in these patients included skin lesions, meningitis, otitis, and shoulder joint arthritis. S. marcescens had not been isolated from this hospital previously, and there were no other S. marcescens infections on other wards of the same hospital or in other hospitals in Jerusalem. On inspection, it was eventually found that a bottle of 5% glucose in saline that had been administered to children on the ward was contaminated with S. marcescens. After the solution was discarded, there were no more S. marcescens cases at that hospital (314). A case of S. marcescens endocarditis occurred in 1953 in a 38-year-old patient from the former Gold Coast, now Ghana. The patient was treated with chloramphenicol and streptomycin but eventually died. S. marcescens was recovered twice from blood cultures and also from postmortem vegetation material (177). In 1957, Gale and Lord reported a case of apparent hemoptysis caused by S. marcescens. The patient, a 39-year-old veteran, had been coughing up red sputum in 1953, and S. marcescens was recovered from the sputum (139). The patient was probably not truly ill with S. marcescens. This case is very similar to the case described by Woodward and Clarke in 1913. S. marcescens was probably the causative agent of a case of empyema in a 55-year-old male patient in Greece with a right spontaneous pneumothorax in 1957. The patient recovered after chloramphenicol treatment (294). In addition, Robinson and Woolley described a case of pseudohemoptysis caused by S. marcescens in 1957 (324). In 1960, Bernard and others described a case of S. marcescens pneumonia in a 33-year-old female patient who had a tubo-ovarian abscess operated on 5 days before symptoms appeared. Penicillin-sensitive Staphylococcus aureus was isolated from abscess material, and the patient was discharged before she developed pneumonia. The patient’s sputum was red, and this was felt by the authors to be due to S. marcescens pigmentation. S. aureus was also isolated repeatedly from sputum

766

MAHLEN

specimens from the patient. The patient was given penicillin, chloramphenicol, and kanamycin over her hospital stay of 58 days, and she eventually recovered; S. marcescens was recovered from 31 sputum cultures over this time (34). Thus, by the end of the 1950s, several cases of infection in humans due to S. marcescens had been described (Table 2). Even so, the belief that S. marcescens was a mostly harmless saprophyte persisted. The fact that the organism can be a pathogen under the right circumstances has been seen a great number of times, though, particularly in nosocomial outbreaks and other opportunistic infections. Opportunistic infections caused by S. marcescens. Initial documented cases revealed the pathogenic potential of S. marcescens. Several of these infections due to S. marcescens were probably hospital acquired in origin, and this bacterium has often been isolated from nosocomial infections or from patients with underlying medical problems. Since S. marcescens is often involved in nosocomial infections, one of the dangers associated with the organism is the potential of intrahospital spread and outbreaks. The first paper that described a series of opportunistic infections caused by S. marcescens was the report by Wheat and others that described 11 cases of S. marcescens UTI, all in adult patients that were immunocompromised to some degree and had indwelling catheters (407). The source of the organism was not clear, and the involved strains were not typed. Wheat and others theorized that risk factors included the indwelling medical devices, the fact that the patients had been ill, and the increased use of antibiotics that may have enabled a normally saprophytic organism to cause disease (407). The next report of a series of nosocomial infections attributed to S. marcescens was the outbreak attributed to contaminated intravenous solutions in a newborn nursery reported by Rabinowitz and Schiffrin in 1952. This was the first reported series of nosocomial infections where a reservoir of S. marcescens was found (314). These two case series are fairly typical accounts of S. marcescens nosocomial outbreaks or clusters of opportunistic infections. Since the early 1950s, there have been a large number of described outbreaks among both adult and pediatric patient populations. (i) Opportunistic infections in adult patients. After Wheat et al. described the UTI cases in San Francisco in 1951, the next case series of human infections due to S. marcescens was published in 1962 by Gale and Sonnenwirth. During a 6-month period from late 1958 to 1959 at Jewish Hospital, St. Louis, MO, nine patients had infections due to S. marcescens. Twelve isolates were recovered from the patients, from wound specimens, empyema drainage, urine, and a throat culture. All of the patients acquired S. marcescens during their hospital stay, and all but one of the patients had been treated with antibiotics prior to infection with S. marcescens. This information led Gale and Sonnenwirth to theorize, like Wheat and others, that increased antibiotic therapy may enable organisms that are normally not pathogens, such as S. marcescens, to cause disease in compromised patients. Eight of the strains were typed at the CDC. The O antigens were type 5 for all strains, while the H antigens of five strains were type 13, that of one strain was type 11, and those of two of the strains were related to both types 11 and 13. Since variability may have been present in H types 11

CLIN. MICROBIOL. REV.

and 13, all of the strains may have been related (140). Several cases of UTI occurred at the University of Washington hospital around the same time frame, between 1959 and 1961. Fourteen symptomatic cases of UTI and four probable cases of S. marcescens UTI occurred in seriously ill, catheterized patients. S. marcescens was recovered from the urine of two other patients without apparent infection. Eight of the isolates were typed at the CDC; only two of the strains had the same type, so this was probably not an outbreak due to a single S. marcescens strain (231). Another series of UTIs caused by S. marcescens was described by Taylor and Keane in 1962. A patient with a chronic UTI was transferred to the Manchester Royal Infirmary from another hospital, and S. marcescens was isolated from his urine. Within a month, six other patients on the same ward had S. marcescens UTIs. Each of the patients were catheterized, leading the authors to suppose that catheterization was a risk factor for S. marcescens infection. The S. marcescens strains were pigmented at room temperature but not when they were incubated at 37°C (376). Other than biochemical characterization, no strain typing was performed. During a 1-year period from 1963 to 1964, 181 isolates of S. marcescens were recovered from specimens collected from 104 patients at the Yale-New Haven Hospital, New Haven, CT. Of particular interest, only one of the isolates was pigmented. Strains were isolated evenly from clinical specimens throughout the year, and 17 of the isolates were serologically typed at the CDC. Sixteen of the isolates had the same type (O9:H5). All of the patients had an underlying illness, an operation, or both. Most (⬃80%) of the patients had received antibiotic therapy before infection with S. marcescens occurred. Clinical specimens from which S. marcescens was isolated included urine, wound specimens, respiratory tract specimens, stool, and blood. The organism was not recovered from environmental sampling in the hospital or from respiratory equipment (81). Dodson described 16 cases of septicemia due to S. marcescens that occurred from 1961 to 1966 at two different hospitals in Birmingham, AL. All of the patients had an underlying disorder, and 13 had received antibiotics prior to septicemia caused by S. marcescens. Nine of the patients died, and S. marcescens was recovered from specimens other than blood, including sputum and urine, for most patients. The respiratory tract was thought to be a portal of entry for three of the patients, and the genitourinary tract was suspected for four patients who had indwelling bladder catheters. Six of the patients that died had received corticosteroids during therapy, prompting Dodson to conclude that this may have been a risk factor. Several of the S. marcescens isolates were not pigmented (108). Eighty-four pigmented S. marcescens isolates were recovered from 49 different patients during a 5-month period from 1967 to 1968 from the same hospital in Columbus, OH. All but one of the patients were adults. S. marcescens isolates were recovered from sputum, urine, various wounds, blood cultures, and stool. Rigorous environmental testing was performed in the hospital, and S. marcescens was recovered from several intermittent positive-pressure breathing machines, from vials of saline used to prepare injectable medications, and from jugs of saline used to irrigate catheters and wounds. Serologic typ-

VOL. 24, 2011

ing was performed at the CDC for some of the patient and environmental isolates, and they were found to be of the same type (58). Since the late 1960s, a tremendous number of nosocomial outbreaks attributed to S. marcescens have been described for adult patients; Farmer and others noted that by 1977, more than 100 outbreaks due to S. marcescens had been described (129). Outbreaks have occurred in medical wards and medical ICUs (112, 147, 280, 320, 329, 335, 383, 408), a hepatologic intensive care unit (306), various surgery units and wards, including cardiac, urology, and neurosurgery wards (17, 19, 43, 102, 103, 106, 113, 121, 124, 186, 202, 213, 237, 289, 293, 299, 304, 319, 327, 349, 360, 389, 390, 397, 409, 418), dialysis units (223), obstetric wards (365), bone marrow transplant and oncology units (221), a pulmonary ward (391), a gastrointestinal disease ward (382), neurology wards (242, 349), and an outpatient pain clinic (83). In some nosocomial outbreaks, S. marcescens was isolated from patients from wards and units throughout hospitals (53, 79, 87, 93, 120, 178, 196, 246, 247, 284, 287, 337, 339, 340, 369, 378, 379). On more extreme occasions, S. marcescens strains have been isolated from outbreaks from more than one hospital in a city or area (53, 93, 155, 183, 202, 284, 339, 340). In other incidents, S. marcescens nosocomial outbreaks occurred in multiple hospitals in the same city or area, but whether the same strain was involved in all of the hospitals is not clear because typing was not performed (247). (a) Multistate outbreaks. Recently, multistate outbreaks of bloodstream infection due to S. marcescens have made headlines. In 2005, two separate outbreaks of S. marcescens bloodstream infections were brought to the attention of the CDC, and both were linked to contaminated intravenous magnesium sulfate solutions obtained from a national distributor. The first outbreak occurred in Los Angeles, CA, in January 2005 and involved six patients, all of whom had received intravenous magnesium sulfate and subsequently developed S. marcescens bloodstream infections. All six of the S. marcescens isolates had identical pulsed-field gel electrophoresis (PFGE) strain typing profiles. The other outbreak occurred in March 2005 in New Jersey and involved five patients. As with the Los Angeles outbreak, all of the patients developed S. marcescens bloodstream infections after receiving intravenous magnesium sulfate; again, the isolates had the same PFGE profiles, and the New Jersey and Los Angeles isolates were identical. The same S. marcescens isolate was recovered from unopened bags of magnesium sulfate from the same lot. The outbreak officially lasted from 5 January through 26 March 2005, involved 18 total patients, and occurred in three other states besides New Jersey and California (3 cases in North Carolina, 2 cases in New York, and 2 cases in Massachusetts). None of the patients died of S. marcescens infection. The magnesium sulfate was produced by a compounding pharmacy; this is significant because compounded pharmaceuticals are held to different regulatory standards than manufactured pharmaceuticals. It is possible that the source of contaminating S. marcescens in this case was human hands (372). Another notable multistate outbreak of bloodstream infection caused by S. marcescens was due to contaminated prefilled heparin syringes (38, 354, 370). The outbreak occurred initially from November to December 2007 in Texas, and eventually

SERRATIA INFECTIONS

767

nine states were involved, through February 2008 (38, 370). The U.S. Food and Drug Administration inspected the company responsible for preparing the heparin syringes and found that it did not comply with regulatory standards (38). The prefilled heparin was a manufactured pharmaceutical, not a compounded one (38). There were 162 reported bloodstream infections caused by S. marcescens due to prefilled heparin syringes from that particular manufacturer, and four of the patients died (38). The outbreak did not stop immediately when the heparin syringe product was recalled (38, 67, 370). This may have been due to contaminated heparin still present in intravenous catheters that was later flushed; however, prefilled saline syringes produced by the same company were also contaminated with S. marcescens, and when these were also recalled the outbreak ended (67). Another outbreak of S. marcescens bloodstream infections was described for September 2009 in China, where multidose heparin vials were contaminated; this was not due to the same manufactured product that was responsible for the multistate outbreak in the United States (241). In this outbreak, nine patients were affected (241). (b) Sources of outbreaks. Opportunistic infections attributed to S. marcescens have been traced to many different sources over the years. Contaminated ultrasonic nebulizers (320), ventilator nebulizers (374), inhalation therapy medications (335), inhalation therapy stock solutions (391), air conditioning units (223, 304), shaving brushes used prior to surgery (237, 408, 409), pressure transducers (30, 112, 397), tap water from pressure-monitoring equipment (327), urine-measuring containers, urinometers, urine-collecting basins, and urinals (147, 329, 349, 356, 418), a cystoscopy area (222), sinks (202, 356), bronchoscopes (304, 353, 389), reusable rectal balloons (61), electrocardiogram leads (360), vitrectomy apparatuses (211), theater linen (124), glass syringes used for preparing intravenous injection fluids (382), saline solutions (66), heparinized saline (375), cream used for obstetric pelvic examinations (365), liquid nonmedicated soap (337), a liquid soap dispenser (374), a finger ring (201), tap water used to take oral medications (186), betamethasone injections (77), an anesthetic (propofol) (33, 181, 278), a narcotic (fentanyl) (289), and transfusion products (315, 342, 403) have all been found to be reservoirs for S. marcescens. Outbreaks associated with either asymptomatic colonized patients or an index, symptomatic colonized patient have occurred several times as well, in some cases including patients colonized in the gut with S. marcescens (19, 130, 206, 221, 356). In addition, many outbreaks are attributed to spread by health care workers (106, 112, 178, 196, 201, 280, 289, 299, 304, 339, 340, 390, 409). S. marcescens contamination of disinfectant solutions, including chlorhexidine, benzalkonium chloride, and hexetidine, has been affiliated with hospital outbreaks as well (43, 120, 251, 282, 283, 304, 395). Pseudo-outbreaks due to S. marcescens have also been described. In some cases, these have been due to contaminated bronchoscopes, resulting in false-positive culture results from respiratory specimens sent to the laboratory (353, 355). In another case, S. marcescens-contaminated EDTA blood-collecting tubes were linked to a pseudo-outbreak of S. marcescens bloodstream infections (130). (c) Typing methods used in outbreaks. Various typing methods were utilized to study strains from several outbreaks. In the

768

MAHLEN

1960s and 1970s, serological typing was the primary method used to determine strain relatedness, in addition to phenotypic characteristics and antibiogram similarity. PFGE has been used in many investigations and is a very reliable typing method for Serratia outbreaks (26, 77, 83, 113, 181, 183, 201, 211, 246, 289, 304, 349, 369, 374, 382, 391, 395). Enzyme electrophoresis was used to study isolates in at least one study (155). In more recent years, PCR-based typing methods have been used to study the relatedness of S. marcescens strains from outbreaks. Repetitive intergenic PCR was used by Liu and others to study an outbreak in a neurology ward (242). Random amplified polymorphic DNA PCR (RAPD-PCR) has also been used (43, 102, 106, 196, 211, 284, 293, 390), as well as amplified fragment length polymorphism (AFLP) analysis (103). One study targeted the flagellin gene of S. marcescens for PCR-restriction fragment length polymorphism (PCRRFLP) analysis (297). The importance of fingerprinting strains has been shown by some circumstances where more than one S. marcescens strain was involved in outbreaks or where other S. marcescens strains that were not part of an outbreak were isolated from patients in the same hospital (102, 246, 304, 369, 390, 391). (d) Outbreak risk factors. Certain risk factors have shown up time and again in the large number of outbreaks due to S. marcescens that have been described for adult patients since the 1950s. Extended hospital stay, prolonged use of antibiotics in inpatients, improper infection control practices by health care workers, immune compromise or underlying medical illnesses, and the use of indwelling medical devices such as catheters are all risk factors. S. marcescens strains that have been involved in outbreaks have often been resistant to multiple antibiotics, and this has served to exacerbate infections in hospital settings. (ii) Opportunistic infections among pediatric patients. The outbreak described by Rabinowitz and Schiffrin in 1952 was important in that it was the first outbreak reported for a pediatric population and was also the first outbreak that was traced to a point source, contaminated intravenous solutions (314). In 1966, Stenderup et al. described another case series of S. marcescens-related nosocomial infections from Aarhus, Denmark. Thirteen premature infants from the same hospital ward were all infected with the same nonpigmented S. marcescens strain from February 1964 to June 1965. Seven of the infants developed septicemia, and six died. The other six infants had purulent conjunctivitis, and all recovered. A source of the organism was not identified (364). Also in 1966, McCormack and Kunin described another set of infections in newborns in a nursery. S. marcescens was recovered from five newborns with UTI and from one newborn each with balanitis, omphalitis, and an upper respiratory tract infection. These infections occurred over a period of 3 months at the University of Virginia Hospital in Charlottsville, VA, and prompted a study of the rate of S. marcescens colonization of newborns there. S. marcescens was found colonizing the umbilical tract in 64.5% of babies. The likely source was thought to be contaminated saline (259). Since then, a large number of pediatrics-related outbreaks have been described, and most were reported from the 1980s on. Outbreaks have been noted in neonatal and pediatric ICUs (4, 14, 18, 21, 28, 41, 60, 63, 74, 76, 88, 94, 95, 116, 133, 137, 150,

CLIN. MICROBIOL. REV.

198, 204, 215, 228, 239, 249, 250, 269, 270, 275, 309, 313, 338, 366, 393, 396, 400, 423), neonatal nurseries/units and special care baby units (7, 100, 156, 190, 238, 275, 310, 359, 362, 387, 423), pediatric oncology units (258), and maternity wards/hospitals (35, 48). Outbreaks of sepsis/bacteremia (4, 18, 74, 88, 116, 157, 215, 238, 258, 310, 341, 359, 362, 423), meningitis (74, 88, 116, 157, 362, 423), conjunctivitis (74, 88, 116), UTIs (116), respiratory tract infections (74, 88, 116, 285, 359), and wound infections (362) due to S. marcescens have all been described for pediatric patients since the series of infections described by McCormack and Kunin in 1966 (259). Conjunctivitis appears to be more common in pediatric population outbreaks in hospitals than in adult populations. (a) Sources of outbreaks. From these pediatric nosocomial infection studies, many environmental sources or point sources have been found as reservoirs for S. marcescens, including hands of health care workers and exposure to health care workers (14, 156, 198, 249, 267, 362, 393, 396, 423), contaminated breast milk, formula, and breast pumps (133, 156, 204, 274, 393), contaminated parenteral nutrition (18), an infected neonate as the index patient or colonization of hospitalized infants (28, 63, 100, 148, 238, 269, 270, 275, 338, 362, 400), equipment such as incubators (28, 198), laryngoscopes (95, 204), suction tubes, soap dispensers (52), and waste jars (393), air conditioning ducts (387), contaminated hand brushes (7), contaminated disinfectants and soap (14, 52, 76, 258, 313, 396), cotton wool pads (137), multidose nebulizer dropper bottles (215), and multidose medications (133). (b) Typing methods used in outbreaks, as well as risk factors. As in outbreaks that have occurred in adults, genotyping methods have been used in many pediatric outbreaks to type the involved S. marcescens strains, including sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) analysis of disrupted S. marcescens cells (116), plasmid profiling (18, 41, 157, 258), PFGE (52, 190, 228, 239, 269, 270, 309, 313, 338, 341, 366, 396), ribotyping (150), rep-PCR (239, 393), RAPD-PCR (18), and PCR fingerprinting (366). Voelz and others performed a systematic analysis of several pediatric S. marcescens outbreak studies from 1984 to 2010 that utilized typing procedures to determine clonality. They concluded that either PFGE or PCR-based fingerprinting typing methods were valuable for control of outbreaks. Voelz and others also determined that two or more nosocomially related inpatient S. marcescens cases signals a potential outbreak that should be investigated. In addition, they determined that the following precautions should be followed if an S. marcescens outbreak is suspected: patients should be isolated, barrier precautions should be utilized, antibiotic therapy should be guided by susceptibility testing and not empirically, and environmental sampling for S. marcescens should be performed only if the institution of barrier precautions does not contain the outbreak (398). Voelz and others determined that risk factors for S. marcescens outbreaks include exposure to hands of health care workers, length of hospital stay, and use of antibiotics that may eliminate the normal flora of a patient, similar to those often ascertained for outbreaks that have occurred among adults in hospitals (94, 137, 398). Ocular infections caused by S. marcescens. Infections of the eye are an area where S. marcescens stands out as a pathogen.

VOL. 24, 2011

The organism commonly causes hospital-acquired eye infections (particularly in neonates and children) or disease in previously injured eyes of patients; for example, Samonis and others recently reported that ocular infections due to S. marcescens were the second most common cause of Serratia infections at the University Hospital of Heraklion, Crete, from 2004 to 2009 (333). The organism can, however, also cause eye infections in individuals without eye trauma or an underlying illness. Cases of conjunctivitis, keratoconjunctivitis, endophthalmitis, corneal ulcers, and keratitis due to S. marcescens have been described. Since S. marcescens is a common environmental organism found in water, soil, and other niches, it is well placed for causing eye infections. The first reported S. marcescens ocular infections of humans occurred among the nosocomial series of infections in premature newborns described by Stenderup et al. in 1966. Six cases of purulent conjunctivitis due to S. marcescens were noted. S. marcescens was the only organism isolated from eye secretions in four of the infants, while S. marcescens was mixed with other organisms in the other two cases. The isolates in these cases were nonpigmented and had the same phenotypic profile, but a common source was not identified (364). In 1970, Atlee and others described two cases of keratoconjunctivitis caused by S. marcescens in Portland, OR. The first patient was a 32-year-old female who was badly burned in a housefire. She developed keratoconjunctivitis a week later, and S. marcescens and S. aureus were cultured from purulent eye discharge; the S. marcescens isolate was nonpigmented. The patient did not have previous eye trauma or infection. S. marcescens was recovered from purulent chest, thigh, and cheek lesions over the next 4 weeks, and she eventually died. The second patient was an 82-year-old male with a history of 8 years of bilateral surgical aphakia. After surgery, the patient had gradual bilateral vision loss with scarring and a loss of tear formation. The patient then developed keratoconjunctivitis due to a nonpigmented S. marcescens strain. Initial treatment with topical chloramphenicol was unsuccessful, and the patient was given topical neomycin-polymyxin B-dexamethasone. The patient worsened and was given a 4-week course of topical gentamicin, chloramphenicol, and neomycin and systemic ampicillin. S. marcescens was cultured during the whole course of treatment. Eye patching, eye expression, and artificial tears were utilized as treatments in addition to antibiotic therapy, and the infection eventually cleared (22). Lazachek and others described a corneal abscess caused by S. marcescens in a 9-year-old girl in 1971 after she was struck in the right eye with a fish hook. The girl was healthy with no underlying medical conditions when the accident occurred, and the source of the organism in this case was probably environmental (234). Eye infections caused by S. marcescens are also associated with the use of contact lenses. S. marcescens was the pathogen in 18% to 23% of cases of keratitis in contact lens wearers in two studies (3, 70). In these two studies, S. marcescens was tied with Pseudomonas aeruginosa as the most common cause of eye infections in contact lens wearers (3, 70). In another study, S. marcescens was the most common bacterial isolate from both corneal scrapings and contact lenses obtained from patients with keratitis (99). While P. aeruginosa is probably the

SERRATIA INFECTIONS

769

most common Gram-negative bacterium associated with ocular infections, S. marcescens is also an important eye pathogen, especially among individuals who wear contact lenses. As shown here, S. marcescens has had a long history as a pathogen, and most described cases of human infection due to this organism are nosocomial in origin. While it is the most commonly recovered member of the genus Serratia from human clinical specimens, it is not the only Serratia species capable of causing illness in humans. Several other Serratia species have been identified in human infections, including S. liquefaciens, S. ficaria, S. fonticola, S. odorifera, S. plymuthica, S. quinivorans, and S. rubidaea. S. grimesii has been recovered from human specimens but has not definitively been found as a human pathogen yet. Table 3 presents a summary of instances when Serratia species other than S. marcescens were recovered from human clinical specimens. S. liquefaciens Although S. liquefaciens is isolated infrequently from human clinical samples, it is considered the second most commonly isolated Serratia species; in a study by Grimont and Grimont, S. liquefaciens was isolated from 2% of 1,108 hospitalized patients in France (160). Determining past infections due to S. liquefaciens is complicated, since prior to 1982, this organism was classified as part of the S. liquefaciens complex, along with S. proteamaculans and S. grimesii (163). Because of this, in some cases human infections were reported as caused by the S. liquefaciens complex and were not identified to the species level, or infections by S. proteamaculans or S. grimesii may have been thought to be S. liquefaciens. Like S. marcescens, S. liquefaciens is an environmental organism that has been associated with infections from contaminated medical devices, products, and equipment, including a vitamin C infusion (123), pressure-monitoring equipment (175), neonatal enteral feeding tubes (193), Alsever’s solution (252), and endoscopes (261). There have also been several described instances of contaminated blood products with associated transfusion reactions in humans (44, 117, 171, 200, 326, 412). One of the first documented reports of S. liquefaciens isolates from humans was published in 1971, when 21 isolates were recovered from various respiratory, urine, wound, and ulcer clinical specimens. Of these 21 isolates, 6 were thought to be involved in infection, 15 were felt to be commensals, and most were isolated from mixed cultures. Of the six S. liquefaciens isolates involved in infection, one was isolated from a fatal case of mucopurulent bronchitis, one was from a case of cellulitis, one was from a gangrenous toe ulcer, and one was isolated from sputum from a case of pneumonia (404). In 1973, Ewing and others described 24 human isolates of S. liquefaciens that had been sent to the CDC between 1957 and 1972 (126). The isolates came from a variety of sites, including blood, several respiratory sources, urine, bile, and feces (126). The authors did not discuss whether any of the isolates were involved in infections. Since that paper was written, several other studies have been published describing the isolation of S. liquefaciens from human specimens, and the clinical significance of these isolates is not known (50, 131, 203). Another early reported case of S. liquefaciens infection in a human was described in 1977, when a patient who wore soft

770

MAHLEN

CLIN. MICROBIOL. REV.

TABLE 3. Summary of infections caused by Serratia species other than S. marcescens Organism

Specimen(s)

Comments (references)

S. liquefaciens Blood, urine, central nervous system specimens, respiratory sites, wounds

Second most common Serratia species involved in human infections (160); also involved in outbreaks (115, 132, 171, 344) and infections with contaminated medical equipment and products (44, 117, 123, 171, 175, 193, 200, 252, 261, 326, 412); like S. marcescens, involved in infections at nearly all sites (6, 15, 50, 75, 90, 115, 123, 126, 131, 132, 171, 174, 203, 262, 266, 271, 276, 308, 326, 332, 336, 344, 361, 401, 404, 412)

S. ficaria

Patient with upper respiratory tract infection; patient may have been colonized after eating figs (149) Patient regularly ate figs; organism recovered with 3 other Gram-negative rods (307) Probably a colonizer; no fig association; recovered from two different patients (51) Probably a colonizer (98) Probably a colonizer (98) Four patients infected; gastrointestinal tract was thought to be source for the patients (8, 98) Patient with sepsis; source was probably the gut (98) Patient with endophthalmitis; patient routinely ate figs, but it is unknown if this was source (25) Patient developed cutaneous abscess (97)

Sputum Leg ulcer Respiratory sites Respiratory secretions Knee wound Gallbladder empyema purulence Blood culture Eye Forearm bite site purulence and blood cultures

S. fonticola

Wound and respiratory tract Leg abscess purulence, blood cultures Right hand wound Stool Scalp wound Synovial fluid

S. grimesii

Several sites Blood cultures Several sites?

S. odorifera

Several sites Primarily respiratory tract specimens Blood cultures, probably other sites Blood cultures, urine Sputum Wound cultures Blood cultures Blood cultures Sputum Urine Blood cultures, urine Sputum, blood cultures

S. plymuthica Face wound Respiratory tract Blood cultures, catheter tip Femur wound Blood cultures Wound cultures Peritoneal fluid Blood cultures Blood cultures Peritoneal fluid Right leg wound culture Left femur wound culture S. quinivorans Bronchial aspirates, a pleural effusion sample, blood cultures S. rubidaea

Several sites

Unknown clinical significance, several isolates (131) After patient had car accident (39) After patient had car accident (305) From an immunocompromised patient with diarrhea (154) Recovered from a hunter after he was bitten by a grizzly bear; recovered with several other bacteria (225) Patient with right knee hemarthrosis after falling off bike into hawthorns (154) Third most common Serratia species recovered from human clinical specimens according to one study by Grimont and Grimont (160) Recovered from three patients, but the clinical significance is not clear (131) Nine strains recovered from human specimens and one from a brain abscess, but the clinical significance is not discussed (368) 23 strains isolated from human specimens, but clinical significance is not known (165) 22 biogroup 1 isolates, most of which were probably not pathogenic (131) 27 biogroup 2 isolates, most of which were felt to be pathogenic; 1 isolate recovered from a blood culture from a fatal case (131) Patient with cirrhosis and septic shock (71) Acquired nosocomial infection of patient with pulmonary vascular congestion and bilateral pleural effusion (265); biogroup 1 Surveillance cultures from 2 patients in a cardiothoracic surgery unit; both cultures were biogroup 2 (331) Outbreak of sepsis in 8 infants, due to biogroup 1; probably acquired from contaminated parenteral nutrition fluid; all of the infants died (136) Patient with catheter-related sepsis caused by biogroup 1 (152) Patient with bronchial infection due to biogroup 1 (64) Five patients with UTI (263) Fatal sepsis caused by biogroup 1 in a patient with chronic renal failure and diabetes (89) Patient with pneumonia and sepsis caused by biogroup 1 (235) Patient with burn wound, may have acquired organism from a radiator; not thought to be a pathogen (78) Recovered from 5 different patients; no isolates thought to be pathogenic (131) Patient with sepsis (189) Patient with femur fracture who developed wound infection and osteomyelitis (424) Three patients with sepsis (62) Two patients with surgical wounds (62) Patient with abdominal infection (62) Patient with sepsis and community-acquired pneumonia (317) Patient with rectorrhagia and septic shock (111) Patient with peritonitis undergoing peritoneal dialysis (286) Patient with right leg necrotic cellulitis (298) Patient with left femur fracture (277) Patient with respiratory distress and pneumonia (40)

4th most common Serratia species recovered from human specimens according to study by Grimont and Grimont (160) Respiratory sites, blood cultures, bile, wound cultures 18 strains sent to CDC; clinical significance is unclear (126) Various sites Several strains, but clinical significance not discussed (131, 161, 203) Left eye Patient with endophthalmitis after penetrating trauma to left eye (207) Blood cultures Patient with bacteremia (332) Blood cultures, bile Patient with bile tract carcinoma (388) Urine Three cases of UTI (263) Blood cultures Patient with sepsis (343)

VOL. 24, 2011

SERRATIA INFECTIONS

contact lenses developed a corneal abscess (90). Contact lens cases were found to be contaminated with S. liquefaciens and S. plymuthica in one study (266) and with S. liquefaciens and Pseudomonas aeruginosa in another (336). In the latter study, S. liquefaciens and P. aeruginosa were also recovered from the contact lenses of the patient, and the patient had developed red eye (336). S. liquefaciens has since been found as a cause of eye infections in a few instances (90, 308). S. liquefaciens has been found as the cause of hospital-acquired outbreaks as well. From 1976 to 1982, six S. liquefaciens strains were recovered from infants in a neonatal nursery in East Melbourne, Australia. Three of the infants had lifethreatening infections caused by S. liquefaciens; the organism was isolated from blood and CSF in one case and from blood in the other two cases. All three of the neonates survived after appropriate treatment (132). In 1984, Serruys-Schoutens and others described a nosocomial outbreak in Belgium involving 10 urinary tract infections due to S. liquefaciens that occurred in about a 3-month period. Each of the patients developed a urinary tract infection with the organism after cystometry or cystoscopy. S. liquefaciens was isolated from the fluid inside the disposable dome of the cystometer, and the outbreak stopped when the dome was replaced as it should have been. All of the patients recovered uneventfully (344). In addition, Dubouix and others described an outbreak of S. liquefaciens among neurosurgery patients in 2005. The organism was isolated from a total of 17 hospitalized patients, primarily from respiratory secretions, but also from urine, a wound, and cerebrospinal fluid. Two of the patients developed sepsis (115). Probably the most publicized outbreak involving S. liquefaciens occurred at a hemodialysis center in Colorado. Ten S. liquefaciens bloodstream infections and six pyrogenic reactions (with no bloodstream infection) occurred within a month in 1999 among outpatients at the center, and all but one of the infections occurred in one section of the dialysis center. The dialysis center had pooled single-use vials of epoetin alfa and then administered the drug to the patients. S. liquefaciens was recovered from pooled epoetin alfa and from empty vials and, additionally, was found in antibacterial soap and hand lotion. All of the S. liquefaciens isolates were identical by PFGE, and the outbreak stopped when pooling of epoetin alfa was discontinued and the soap and lotion were replaced. All of the patients recovered with antimicrobial therapy (171). There have been several other published case reports involving S. liquefaciens as a human pathogen. The organism has been isolated as a cause of abscesses (361), endocarditis (75, 276), a fistulous pyoderma (401), fatal meningoencephalitis (15), septic arthritis (174), septicemia (6, 115, 123, 132, 171, 326, 332, 412), and urinary tract infections (263, 344) and from a wound culture after a man received a swordfish bill injury (262). S. ficaria There have been several instances of S. ficaria reported as a causative agent of disease in humans, many of which had a link to figs. The first reported isolation of S. ficaria from a human specimen was in 1979, when it was isolated from the sputum of a patient with an upper respiratory tract infection. S. ficaria was isolated from the patient’s sputum a day or two after she

771

had eaten a fig, and it was thought that the isolate was probably a transient upper respiratory tract or mouth colonizer (149). S. ficaria was isolated from a leg ulcer from a patient in Hawaii in 1980, along with three other Gram-negative rods (307). This isolate was considered to have contributed to disease; it is notable that this patient regularly ate prunes. Pien and Farmer also reported that S. ficaria was identified retroactively after being isolated from the nasogastric tube from a patient in Hawaii in 1977, although no other clinical information is available (307). In 1982, S. ficaria was cultured from the respiratory specimens of two different patients in Hornu, Belgium. In both cases, S. ficaria was felt to be a colonizer. Apparently neither patient had consumed figs, and the source of S. ficaria from both patients is not known (51). In the 1980s and 1990s, S. ficaria was isolated several times from human specimens in France. The organism was recovered from respiratory secretions from a patient in 1983 and from a knee wound culture in 1988; in both cases, S. ficaria was thought be a colonizer and a nonpathogen (98). S. ficaria was isolated as the cause of infection four different times during the 1990s for purulence from patients with gallbladder empyemas (8, 98). One of the patients had regularly eaten figs, but apparently the timing did not coincide with infection (8). The source of the organism in each of these cases was probably the gut of each patient, so S. ficaria may also colonize the human gastrointestinal tract (8, 98). Each of the patients was considered to be immunocompromised prior to infection (8, 98). S. ficaria was also recovered from blood from a patient in France with adenocarcinoma of the pyloric antrum who developed septicemia, and this was also thought to be a true infection; again, the patient was immunocompromised (98). The source of S. ficaria in this case was also the gut of the patient (98). All of the gallbladder empyema patients and the patient with sepsis responded well to therapy. In 2002, Badenoch and others reported a case of endophthalmitis caused by S. ficaria in a 73-year-old man in Australia. The infection resulted in the loss of the patient’s eye. The patient evidently had eaten figs for a large part of his life, but the source of the organism that was recovered from eye cultures is not known. S. ficaria could have been a well-established member of the patient’s flora by the time the eye infection occurred. The patient had a history of previous eye trauma, so combining this with his age, he was considered to be immunocompromised (25). The last reported human infection caused by S. ficaria occurred in an otherwise healthy 47-year-old man in Greece. The man was a hunter and was bitten by a wild dog on his forearm and his shoulders. A cutaneous abscess developed at the forearm bite site, and S. ficaria was isolated from purulence from the abscess and from blood cultures (97). This human infection is probably the first known infection caused by S. ficaria in a patient who was not compromised in some way and shows the potential of the organism to be involved in zoonotic infections. S. fonticola S. fonticola was first reported from human specimens in 1985, when Farmer and others studied several wound culture and respiratory tract isolates (131). The clinical significance of these isolates is unknown. The first human infection caused by

772

MAHLEN

CLIN. MICROBIOL. REV.

S. fonticola was reported in 1989, when it was recovered in pure culture from leg abscess purulence and from a blood culture bottle from a 73-year-old female patient who had been in a car accident in France (39). In 1991, S. fonticola was isolated as the predominant organism from a right hand infection of a 39year-old woman after she had also been in a car accident (305). S. fonticola was then isolated in 2000 from the stool of an immunosuppressed patient with diarrhea in France (154). S. fonticola was later isolated from scalp wounds of a 49-year-old hunter after he was attacked and bitten by a grizzly bear in Alberta, Canada. S. fonticola was isolated in this case with several other bacteria, including S. marcescens (225). In 2008, S. fonticola was recovered from synovial fluid from a 15-yearold boy with right knee hemarthrosis in France. The boy had fallen off a bike and into hawthorns, so it is likely that S. fonticola was present on the thorns (154). S. grimesii There have been few descriptions of S. grimesii isolated from human specimens. Among the 1,108 Serratia species from hospitalized patients from France that Grimont and Grimont studied, 0.5% were identified as S. grimesii. This ranks S. grimesii as the third most commonly isolated Serratia species in their study (160). Farmer and others studied three isolates from blood cultures from France, but no clinical information is available for these strains (131). Lastly, nine S. grimesii strains from human specimens were described by Stock and others (368). The clinical significance of the strains is not discussed, although one strain was isolated from a brain abscess (368). S. odorifera S. odorifera was first named in 1978 when Grimont and others characterized 25 related strains. Twenty-three of the strains were isolated from human specimens, although clinical significance was not established for any of them. Two different biogroups, 1 and 2, have been identified (165). In 1985, Farmer and others described 22 S. odorifera biogroup 1 isolates and 30 biogroup 2 isolates; 16 of the biogroup 1 isolates were recovered from human specimens, and 27 of the biogroup 2 isolates were from human specimens. The S. odorifera biogroup 1 isolates from this study, most of which were isolated from the respiratory tract, apparently were not actually involved in clinical infections, prompting the authors to doubt the disease potential of biogroup 1 strains. The S. odorifera biogroup 2 isolates from this study were more commonly isolated from specimens, though, suggesting a more invasive source, such as blood cultures, although few clinical data were supplied for the strains. One of the blood culture isolates was from a fatal case, but there is no more information available (131). The first probable case of confirmed human infection caused by S. odorifera was reported in 1988 in Florida for a 67-year-old male with cirrhosis. The patient was a chronic alcoholic and was admitted with septic shock. S. odorifera biogroup 1 was isolated from both blood and urine. Antibiotic therapy with amikacin and cefotaxime cleared the infection (71). The next documented human case involving S. odorifera was a nosocomial infection that occurred in 1990 in Wisconsin in a 73-year-old man admitted with progressive claudication.

The patient had several underlying medical issues, including chronic obstructive pulmonary disorder, chronic renal failure, and severe atherosclerotic vascular disease. The patient developed pulmonary vascular congestion and bilateral pleural effusion while in the hospital, and S. odorifera biogroup 1 was cultured from sputum specimens. The patient was treated empirically with tobramycin, metronidazole, ceftriaxone, and trimethoprim-sulfamethoxazole and recovered with ceftriaxone therapy after the identity and susceptibilities of the organisms were determined. The authors also described that two other S. odorifera biogroup 1 isolates had been recovered at the University of Wisconsin hospital; both of these isolates were recovered from immunocompromised patients. Both isolates were from sputum, and one was also cultured from a catheter tip (265). Nosocomial transmission of S. odorifera has been documented a few more times since 1990. In 1994, S. odorifera biogroup 2 was isolated from surveillance wound cultures from two patients in a cardiothoracic surgery unit at the University of Iowa; the source of S. odorifera in these cases is not known (331). Also in 1994, an extreme outbreak due to S. odorifera was described by Frean and others, when eight infants died of S. odorifera biogroup 1 septicemia because of contaminated infant parenteral nutrition fluid in South Africa. The origin of the contaminated parenteral nutrition fluid was not clear for this outbreak (136). There have been several other instances of S. odorifera infection in humans. In 1994, S. odorifera biogroup 1 was reported as a cause of catheter-related sepsis in a 19-year-old woman. The patient had a history of thalassemia major and had a Broviac catheter placed 2 months prior to this infection (152). A bronchial infection due to S. odorifera biogroup 1 was reported from France in 1999 (64), and five instances of S. odorifera UTI were described from Brazil in 2004 (263). In another case, fatal sepsis caused by S. odorifera biogroup 1 occurred in a 73-year-old woman. This patient had a history of cirrhosis, adult-onset insulin-dependent diabetes mellitus, and idiopathic thrombocytopenic purpura and had a left nephrectomy performed 30 years prior. In addition, the patient had chronic renal failure and was receiving long-term dialysis. S. odorifera was isolated from several blood cultures and a urine culture in this case (89). Lastly, a case of pneumonia and septicemia caused by S. odorifera biogroup 1 was described for a 57-year-old patient with an underlying history of chronic hepatitis C virus infection, alcoholic liver disease, chronic bronchitis, paranoid schizophrenia, and past injection drug use. It is not clear in this case whether the portal of entry in the patient was the lungs or whether the pneumonia was secondary to sepsis (235). S. plymuthica Clark and Janda first reported the isolation of S. plymuthica from a human clinical specimen in 1985, when the organism was recovered from a surveillance culture from a burn wound on the face of an 8-month-old boy. The boy received the burn wound after falling into a steam radiator, and the organism was probably acquired from the radiator. In this case, S. plymuthica was probably not a pathogen (78). In 1985, Farmer and others also described five isolates of S. plymuthica that were isolated

VOL. 24, 2011

from the respiratory tracts of humans; none were from human infections (131). There have been several reported human infections caused by S. plymuthica. The first documented case of S. plymuthica infection in humans occurred in 1986 in Westchester County, NY. S. plymuthica was isolated from blood cultures and a central venous catheter tip culture from a 54-year-old alcoholic man who had previously been diagnosed with cirrhosis. The patient improved with ampicillin, gentamicin, and clindamycin therapy; the isolate was sensitive to gentamicin (189). A second S. plymuthica human infection case occurred in Switzerland in 1987. An 18-year-old patient was admitted with a distal right open femur fracture after a motorcycle accident. The site became infected a few months later, and eventually osteomyelitis developed. S. plymuthica was isolated from the wound site as the predominant organism; gentamicin spherules were added to the operation site after wound excision and drainage, and the patient improved (424). Carrero and others described a series of S. plymuthica isolates recovered from blood cultures (three cases) and surgical wound exudate cultures (two cases), with a sixth isolate recovered from peritoneal fluid; the cases all occurred from 1989 to 1990 in Spain and were serious infections. The sepsis and surgical wound culture cases were probably nosocomial in origin, since all of the patients developed infection at least a few days after admission. All of the patients recovered after therapy with drainage, an aminoglycoside, a broad-spectrum cephalosporin, or a combination of an aminoglycoside and a ␤-lactam antibiotic; however, one patient died due to underlying illness (62). Another case of S. plymuthica sepsis was reported in 1992 for a 50-year-old woman diagnosed with community-acquired bacteremia. The patient presented initially with a 3-day history of dyspnea, a dry cough, and thoracic pain. S. plymuthica was recovered from blood cultures, and the patient was successfully treated with a combination of gentamicin and erythromycin (317). A case of nosocomial sepsis caused by S. plymuthica in a 79-year-old patient was also described in Spain in 1994. The patient was admitted with rectorrhagia and developed septic shock a week after admission; the patient improved with antimicrobial therapy (111). In 2000, S. plymuthica was isolated from a case of peritonitis in a 74-year-old male with continuous ambulatory peritoneal dialysis. The patient was initially treated with gentamicin and vancomycin and did not get better, but he improved after piperacillin was added. The patient, however, died later due to cardiac difficulties (286). S. plymuthica was isolated as a cause of necrotic cellulitis from a 66-year-old female patient in 2003. The patient had steroid-dependent asthma and had initially presented with a right inferior extremity contusion wound. She was admitted 2 weeks later with signs of Cushing’s disease, and her right leg was red with an erythematous erosion present. S. plymuthica was recovered from both blood cultures and from cellulitis cultures. Surgical exploration, debridement, and therapy with imipenem were successful in treating the infection (298). The organism was also involved in a case of septic pseudoarthrosis published in 2008 from a 17-year-old patient with postoperative left thigh pain. The patient had a left femur fracture treated with an osteosynthesis plate 10 months prior to presentation. S. plymuthica was recovered from a swab sample taken from pinkish fungos-

SERRATIA INFECTIONS

773

ities that were observed around two proximal screws at the site. The patient was treated with ciprofloxacin and gentamicin and recovered (277). S. quinivorans The first, and at this time only, human infection caused by S. quinivorans occurred in 1990 in France in a 43-year-old homeless man. The patient was an alcoholic and was admitted with a mouth abscess that eventually caused an obstruction, so a tracheotomy tube was placed. The patient later developed respiratory distress and pneumonia. S. quinivorans was isolated from bronchial aspirates, a pleural effusion sample, and blood cultures. The patient died of multisystem organ failure a little over a month after admission (40). The patient could have acquired the organism while sleeping outside due to being homeless. S. rubidaea While S. rubidaea has been isolated from human specimens, its pathogenic potential in humans appears to be very limited. S. rubidaea was isolated from 0.2% of 1,108 Serratia species from hospitalized patients in France, making it the fourth most common Serratia species identified from human specimens in that study (160). S. rubidaea has been detected in human specimens from several other studies. In 1973, Ewing and others described 18 S. rubidaea strains that were sent to the Centers for Disease Control and Prevention between 1957 and 1972 and had been isolated from the respiratory tract, blood, a few wound specimens, and bile; the authors did not elaborate on whether any of the strains caused infection (126). Since then, S. rubidaea has been isolated from human specimens in several studies from various sources (131, 161, 203). The clinical significance of S. rubidaea was not described in any of these surveys. At this time, there have been only a few published cases of human infection by S. rubidaea. In 1983, Joondeph and Nothnagel described a case of endophthalmitis caused by S. rubidaea in a 10-year-old boy after he had penetrating trauma in his left eye. Treatment with several topical antimicrobial agents cleared the infection (207). A case of S. rubidaea bacteremia in a patient with cancer was published in 1989 by Saito and others. The infection was cleared with antibiotic therapy (332). In 1994, S. rubidaea was isolated from blood and bile from a 64-year-old female patient with bile tract carcinoma in Spain. The patient recovered after antibiotic treatment. It could not be determined where or how the patient acquired S. rubidaea, and it was theorized that the source was endogenous (388). Two other papers since 1996 have described human infections by S. rubidaea; three strains were isolated from urinary tract infections in Brazil, and a case of sepsis was described in Tunisia (263, 343). VIRULENCE FACTORS OF SERRATIA SPECIES Serratia species are usually opportunistic pathogens, and virulence factors produced by these bacteria are not understood well. All of the species in the genus are motile, and quorum sensing (QS) has been described for some of these organisms.

774

MAHLEN

S. marcescens is capable of producing well-known virulence factors such as fimbriae for adherence (24, 135, 357, 415). In 1997, Hejazi and Falkiner wrote a review paper and summarized the virulence factors known at the time (179). The S. marcescens RssAB-FlhDC-ShlBA Pathway S. marcescens produces a hemolysin, ShlA, that functions as a pore-forming toxin in concert with another protein, ShlB; together, these proteins cause cytotoxicity in red blood cells and in other eukaryotic cells, such as epithelial cells and fibroblasts (184, 226). They are contact dependent and are not released as extracellular products (184). Without ShlB, ShlA is inactive. ShlB is an outer membrane protein, and it activates and secretes ShlA. Activation of ShlA is also dependent on phosphatidylethanolamine, a primary component of the S. marcescens outer membrane. In addition to cytotoxic activity, ShlA also mediates release of inflammatory molecules and apparently contributes to uropathogenicity. Since ShlA and ShlB are cell associated, the ability of these proteins to cause damage usually depends on the ability of S. marcescens to adhere to eukaryotic cells (184). S. marcescens has the ability to swarm at 30°C on LuriaBertani agar, and swarming has been shown to be a pathogenic factor for Proteus mirabilis and Pseudomonas aeruginosa (5, 227, 290). At 37°C, a two-component system, RssAB, inhibits swarming and also decreases the production of ShlA (227, 240). Lai and others showed that without the RssAB regulatory system, swarming and hemolytic activities were increased (227). Another system, FlhDC, controls expression of flagella for enteric bacteria and is important for quorum sensing (392). FlhDC also positively regulates production of hemolysin and is involved in swarming and biofilm production in S. marcescens (240, 392). When RssAB is activated, flhDC expression is reduced, ShlA hemolysin is produced, and biofilm formation occurs (240). If RssAB is deleted or nonfunctional, flhDC expression increases, and S. marcescens produces increased amounts of hemolysin, swarms, does not form a biofilm, and becomes more virulent (240). The RssAB-FlhDC-ShlBA pathway appears to be important for pathogenesis of S. marcescens. Quorum Sensing in Serratia Species QS, a cell-to-cell signaling mechanism employed by many bacteria, has been described for S. marcescens, S. plymuthica, and S. proteamaculans (243, 392). Quorum sensing is used by bacteria to control certain biological functions, such as biofilm formation and the production of antibiotics (392). When cell populations reach a critical mass, signaling molecules are released that allow bacteria to respond to their environment. Most Gram-negative bacteria, including the aforementioned Serratia species, utilize N-acylhomoserine lactones (AHLs) as the signaling molecules in quorum sensing. The QS system is composed of a LuxI-type AHL synthase and a LuxR-type AHL receptor (392). Various LuxIR-type QS systems have been described for S. marcescens strains. In strain MG1 (formerly called S. liquefaciens), the SwrI/SwrR system regulates swarming motility, biofilm formation, production of serrawettin, protease, and Slayer protein, and fermentation of butanediol (119). In S. marcescens strain 12, the SmaI/SmaR QS system is most re-

CLIN. MICROBIOL. REV.

lated by sequence to the SwrI/SwrR system from strain MG1 and regulates swarming motility, hemolytic activity, biofilm formation, and production of chitinase and caseinase (91). Another S. marcescens strain that has been studied, SS-1, has demonstrated sliding motility that is flagellum independent and regulated by the SpnI/SpnR quorum sensing system (187). Prodigiosin production is also regulated by the SpnI/SpnR system in strain SS-1 (187). The QS system SmaI/SmaR also regulates prodigiosin production and carbapenem biosynthesis in the unnamed Serratia sp. strain ATCC 39006 (381). Different QS systems have also been described for S. plymuthica strains. Two separate LuxIR-type systems, SplIR and SpsIR, have been identified in the plant pathogen S. plymuthica strain G3 (243). These two QS systems regulate antifungal activity, adhesion, biofilm production, and production of exoenzymes, but not swimming motility, in this strain (243). S. plymuthica strain HRO-C48, also a plant pathogen, has a SplIR QS system that also regulates antifungal activity and production of exoenzymes (279). However, the QS system of strain HRO-C48 does not regulate biofilm production or adhesion and does regulate swimming motility (279). Thus, QS systems may be strain dependent and may reflect the particular environment and/or lifestyle of a given strain. Bacteria that form biofilms are important in medicine because they can colonize catheters and other indwelling devices. In addition, bacteria can form biofilms on contact lenses and contact lens cases, and this has been identified as a risk factor for P. aeruginosa eye infections (316). The production of biofilm may represent the typical environmental form of many bacteria and gives several significant advantages, including increased resistance to antibiotics and the immune system (104, 243). Biofilm production has been reported for several Serratia species, including S. marcescens and S. plymuthica (243, 346). Quorum sensing appears to play a role in regulating biofilm production for Serratia species, as described above. In addition, Shanks and others found that the oxidative stress response transcription factor OxyR plays a role in S. marcescens biofilm formation (346). It is theorized that biofilm production plays an important role in the pathogenesis of S. marcescens, although in one study by Pinna and others, isolates of S. marcescens and S. liquefaciens recovered from soft contact lensrelated corneal ulcer cases did not produce biofilms. Rather, it was thought that exoenzymes produced by S. marcescens and S. liquefaciens may play a role in keratitis (308). Enzymes Produced by Serratia Species While the ShlAB hemolysin of S. marcescens is contact dependent, an extracellular hemolysin was described in 1989 and was recently characterized (153, 351). This hemolysin, PhlA, has phospholipase A activity (351). PhlA does not apparently have direct cytolytic activity; however, it acts upon phospholipid and produces lysophospholipid, which was cytolytic for human, horse, and sheep red blood cells and the HeLa and 5637 cell lines (351). S. marcescens and other Serratia species produce many other enzymes, such as metalloproteases, gelatinase, and alkaline protease, that may enable the organism to cause infections, particularly diseases of the eye (256, 308). Several proteases are described in a review by Matsumoto; the described pro-

VOL. 24, 2011

SERRATIA INFECTIONS

775

TABLE 4. Antibiogram of S. marcescens susceptibilities at three different Army medical facilities, in Pierce County, WA, from two MYSTIC surveys, and from the TEST survey % Susceptibilityh (n) Antibiotic

Madigan Healthcare System (110)a

Pierce County, WA (339)b

Tripler Army Medical Center (138)c

Walter Reed Army Medical Center (29)d

MYSTIC Program European data (195)e

TEST U.S. data (427)f

MYSTIC Program U.S. data (145)g

Amikacin Cefepime Ceftazidime Ceftriaxone Ciprofloxacin Gentamicin Imipenem Levofloxacin Meropenem Piperacillin-tazobactam Tobramycin Trimethoprim-sulfamethoxazole

98 100 100 97 95 98 97 100 100 97 96 100

NR NR 100 98 91 99 98 95 NR 98 97 97

100 100 99 99 94 99 100 98 NR 97 91 98

100 100 100 97 90 100 100 97 NR 95 79 NR

NR NR 93.9 NR 92.3 96.7 99.5 NR 100 88.7 91.5 NR

98.6 96.0 92.3 91.8 NR NR 100 93.7 98.3 95.8 NR NR

NR 97.9 98.6 95.9 91.7 NR 97.2 95.9 97.2 93.8 91.7 NR

a

Combined data for 2008 to 2010. Madigan Healthcare System is located in Tacoma, WA. 2009 data. c Combined data for April 2009 to April 2011. Tripler Army Medical Center is located in Honolulu, HI. d 2010 data. Walter Reed Army Medical Center is located in Washington, DC. e 2007 data on European medical centers from the MYSTIC Program (386). Data are for the following Serratia species: S. marcescens (170 isolates), S. liquefaciens (19 isolates), unidentified Serratia species (3 isolates), S. fonticola (2 isolates), and S. odorifera (1 isolate). f 2007 data on U.S. medical centers from the Tigecycline Evaluation and Surveillance Trial (TEST) (114). g 2008 data on U.S. medical centers from the MYSTIC Program (318). Data are for the following Serratia species: S. marcescens (119 isolates), S. liquefaciens (5 isolates), and unidentified Serratia species (21 isolates). h NR, not reported. b

teases affect defense-related humoral proteins and various types of tissue cells (256). A recently described metalloprotease from S. grimesii, grimelysin, is proteolytic for actin (46). E. coli that expressed grimelysin was able to invade Hep-2 cells, so this metalloprotease may allow bacterial internalization into eukaryotic cells (47). ANTIMICROBIAL RESISTANCE OF SERRATIA SPECIES As with most literature regarding Serratia species, the vast majority of antimicrobial resistance that has been described for this genus has occurred in S. marcescens. The fact that S. marcescens was a very resistant organism was recognized in early published cases. For example, Wheat and others, in their seminal report of 11 cases of UTI from San Francisco in 1951, reported probable resistance of the isolate that caused fatal endocarditis to polymyxin B, terramycin (oxytetracycline), chloramphenicol, streptomycin, and penicillin, with moderate sensitivity to sulfonamides (407). It is now known that S. marcescens is frequently resistant to multiple antibiotics. Outbreaks caused by multiply resistant S. marcescens strains have been described, and many S. marcescens strains carry both chromosomally encoded and plasmid-mediated resistance determinants for several different types of antibiotics. Indeed, one of the hallmarks of nosocomial outbreaks due to S. marcescens is very resistant strains, making such outbreaks even more devastating for compromised patients. Typical Resistance Patterns of Serratia Isolates Like other members of the Enterobacteriaceae, S. marcescens and other Serratia species are intrinsically resistant to penicillin G, the macrolides, clindamycin, linezolid, the glycopeptides,

quinupristin-dalfopristin, and rifampin (244, 367, 368). In addition, most members of the genus Serratia, including S. marcescens, are usually resistant to ampicillin, amoxicillin, amoxicillin-clavulanate, ampicillin-sulbactam, narrow-spectrum cephalosporins, cephamycins, cefuroxime, nitrofurantoin, and colistin (82, 244, 367, 368). If a Serratia isolate tests susceptible to one of these antibiotics, the result should be viewed with suspicion and retested. S. marcescens, S. odorifera, and S. rubidaea were intrinsically resistant to tetracycline in studies by Stock and others (367, 368). S. marcescens also harbors a chromosomal ampC gene that can extend resistance to several more ␤-lactam antibiotics. In addition, some strains carry chromosomally encoded carbapenemases, and plasmid-mediated enzymes can be acquired that further extend resistance to ␤-lactams. Sensitivities to other antimicrobials, such as the quinolones and trimethoprim-sulfamethoxazole, are more variable. In general, most Serratia species are sensitive to the aminoglycosides (367, 368). Sensitivity of S. marcescens strains to aminoglycosides, though, is more variable, and S. marcescens has a chromosomal aminoglycoside resistance gene that may contribute to decreased susceptibility. At my medical facility in Tacoma, WA, most S. marcescens isolates are sensitive to commonly prescribed antimicrobial agents. Antibiogram data for 110 different patient isolates recovered from clinically significant infections are shown in Table 4, compared to data for Pierce County, WA, and two other U.S. Army medical facilities (Tripler Army Medical Center, Honolulu, HI, and Walter Reed Army Medical Center, Washington, DC), 2007 data from European medical centers from the Meropenem Yearly Susceptibility Test Information Collection (MYSTIC) Program (386), 2007 U.S. data from the Tigecycline Evaluation and Surveillance Trial (TEST) (114), and 2008 U.S. data from

776

MAHLEN

the MYSTIC Program (318). The MYSTIC Program antibiograms represent primarily S. marcescens data but also include other Serratia species. The 2007 MYSTIC Program data presented in Table 4 summarize antimicrobial sensitivities for 195 Serratia isolates (S. marcescens, 170 isolates; S. liquefaciens, 19 isolates; unidentified Serratia spp., 3 isolates; S. fonticola, 2 isolates; and S. odorifera, 1 isolate) collected from 28 different European hospitals (386). The 2008 MYSTIC Program data were collected from 15 different U.S. medical centers and show data for 145 Serratia isolates (S. marcescens, 119 isolates; S. liquefaciens, 5 isolates; and unidentified Serratia spp., 21 isolates) (318). Aminoglycoside Resistance in Serratia Aminoglycoside-modifying enzymes are the most common mechanism of aminoglycoside resistance in bacteria. These enzymes modify their targets, aminoglycosides, by adding either an acetyl group (N-acetyltransferases [AAC]), a phosphate group (O-phosphotransferases [APH]), or a nucleotide (O-nucleotidyltransferases [ANT]). The antibiotic then does not bind to the ribosome target. The aminoglycoside-modifying enzymes are usually acquired by bacteria via genes on plasmids. Aminoglycoside resistance in bacteria can also occur because of alteration of the ribosome target, cell impermeability, or efflux. Another type of enzyme, a 16S rRNA methylase called RmtB, has been identified in S. marcescens (110). This enzyme is plasmid mediated and provides high-level resistance to several aminoglycosides, including kanamycin, tobramycin, amikacin, gentamicin, streptomycin, and arbekacin (110). Other plasmid-mediated 16S rRNA methylases have been identified in S. marcescens, including ArmA, RmtA, and RmtC (210). S. marcescens harbors a chromosomal aminoglycoside-modifying enzyme of the AAC(6⬘) family, AAC(6⬘)-Ic (65). Enzymes of the AAC(6⬘)-I class are 6⬘-N-acetyltransferases and are clinically significant in that they may provide resistance to several commonly prescribed aminoglycosides, such as amikacin, tobramycin, and netilmicin. The S. marcescens chromosomally encoded AAC(6⬘)-Ic enzyme is normally expressed weakly or at low levels, and because of this, S. marcescens is normally sensitive to aminoglycosides, and susceptibilities of these antibiotics can be reported. Treatment with amikacin, tobramycin, or netilmicin, though, may result in selection of a hyperproducing mutant of the chromosomal enzyme. In this case, an AAC(6⬘)-Ic-hyperproducing strain will be resistant to amikacin, tobramycin, netilmicin, neomycin, and kanamycin (244, 347). In a survey published in 1985, 19.2% of aminoglycosideresistant Gram-negative rods in the United States were Serratia isolates (350). Of these isolates, 69% carried 6⬘-N-acetyltransferases. Another 18.4% of these Serratia strains carried the ANT(2⬙) enzyme, a 2⬙-O-nucleotidyltransferase; this enzyme confers resistance to gentamicin, tobramycin, and other aminoglycosides. Perhaps more ominously, 47.8% of Serratia strains carried both a 6⬘-N-acetyltransferase and the ANT(2⬙) enzyme, and this combination of determinants confers resistance to nearly all of the clinically useful aminoglycosides. The same survey also found that 42.7% of the examined aminoglycosideresistant Gram-negative rods from Japan, Korea, and Formosa were Serratia isolates. Nearly all of these strains (97.9%) car-

CLIN. MICROBIOL. REV.

ried a 6⬘-N-acetyltransferase, and 71.4% harbored both a 6⬘N-acetyltransferase and the ANT(2⬙) enzyme (350). In another study, antimicrobial sensitivities of a large number of Gram-negative rod isolates that were recovered from ICU patients from hospitals throughout the United States from 1993 to 2004 were examined. S. marcescens was the sixth most commonly isolated organism, representing 5.5% of all Gram-negative rods from the study. Antimicrobial sensitivity data were shown for 2002 to 2004, and 7.1% of the S. marcescens strains were resistant to tobramycin, with 0.8% resistant to amikacin; an additional 5.8% and 1.1% of S. marcescens strains had intermediate resistance to tobramycin and amikacin, respectively (245). Another recent study evaluated amikacin resistance in Enterobacteriaceae isolates from 1995 to 1998 and 2001 to 2006 from a university hospital in South Korea. In this study, 7.5% of S. marcescens isolates were resistant to amikacin, and most of the resistant strains were isolated from 2001 to 2006. Six of the S. marcescens strains carried both ArmA and AAC(6⬘)-1b on plasmids. In this study, there were only four other Serratia species recovered from clinical specimens, and none were resistant to amikacin (210). Many nosocomial outbreaks in both pediatric and adult patients have occurred with S. marcescens strains resistant to one or more aminoglycosides (17, 41, 53, 79, 88, 93, 120, 238, 258, 280, 285, 287, 339, 356, 423). Most of the initial reports of aminoglycoside-resistant S. marcescens nosocomial outbreaks occurred in the mid- to late 1970s (for example, see references 79, 93, and 339). The outbreak described by Craven and others in 1977 is a useful study of probable selection of a hyperproducing aminoglycoside-resistant mutant. Two adjacent hospitals associated with the University of Texas Health Science Center experienced a 22-month nosocomial outbreak of gentamicin-resistant S. marcescens infections. Amikacin was given to 19 patients during this time. Four severely ill patients died within 2 days of being given amikacin; the authors felt that S. marcescens was a key factor in the death of each patient. Ten other patients who were not as ill had S. marcescens infections that responded well to amikacin therapy. S. marcescens infections persisted in the other five patients. In four of these persistent infections, the isolates were initially sensitive to amikacin but became resistant over time. Two of these patients died, one after 7 days of amikacin therapy, and the other after 18 days of amikacin therapy (93). ␤-Lactam Resistance in Serratia Species As already discussed, Serratia species are intrinsically resistant to several ␤-lactam antibiotics, including penicillin G, ampicillin, amoxicillin, amoxicillin-clavulanate, cefuroxime, and narrow-spectrum cephalosporins. All Serratia species are intrinsically sensitive to carbapenems, although some S. marcescens strains have been identified that harbor chromosomal carbapenemases. In addition, most of the members of the genus Serratia carry a chromosomal ampC gene, and there have been several descriptions of strains acquiring plasmidmediated extended-spectrum ␤-lactamases (ESBLs). Chromosomal AmpC ␤-lactamases of Serratia species. AmpC ␤-lactamases are classified as either group 1 enzymes by the Bush scheme or class C enzymes by the Ambler

VOL. 24, 2011

scheme (197). They hydrolyze primarily cephalosporins, including the cephamycins, although these enzymes have activity against the penicillins and aztreonam (197). The chromosomal ampC genes of S. marcescens and several other members of the Enterobacteriaceae are inducible by various ␤-lactam antibiotics by a complex mechanism that involves cell wall recycling (173). The 5⬘-untranslated region (5⬘UTR) of the S. marcescens chromosomal ampC gene was found to be 126 bases long (248). This is longer than those for other Enterobacteriaceae organisms with chromosomal ampC genes. Sequence analysis of the S. marcescens ampC 5⬘-UTR predicted a stem-loop structure that provides stability to S. marcescens ampC mRNA (248). Typically, the expression of AmpC is low from S. marcescens and other members of the Enterobacteriaceae (173, 197). Induction of the chromosomal ampC gene causes an increase in AmpC ␤-lactamase production and increases the MICs of several ␤-lactams (173, 197). Strong inducers of ampC in enteric bacteria such as S. marcescens include cefoxitin, imipenem, ampicillin, amoxicillin, benzylpenicillin, and narrow-spectrum cephalosporins, including cephalothin and cefazolin (197). Broad-spectrum cephalosporins, such as ceftazidime, cefotaxime, and ceftriaxone, and other ␤-lactams, including cefepime, cefuroxime, and aztreonam, are weak inducers (197). Overexpression of AmpC ␤-lactamase in S. marcescens and other Enterobacteriaceae, however, is most often due to a mutation or deletion in the induction/ cell wall recycling pathway (173, 197). These mutants, called derepressed mutants, are clinically important and may result in treatment failures with ␤-lactam antibiotics (197, 244). While S. marcescens and other Serratia species are not intrinsically resistant to broad-spectrum cephalosporins, the use of these antimicrobials in treating Serratia infections is hazardous because the emergence of derepressed ampC mutants occurs more often with these agents than with other antimicrobials (197, 244). The 2011 Clinical and Laboratory Standards Institute (CLSI) Performance Standards for Antimicrobial Susceptibility Testing (M100-S21) (82) includes this warning concerning treatment with broad-spectrum (thirdgeneration) cephalosporins: “Enterobacter, Citrobacter, and Serratia may develop resistance during prolonged therapy with third-generation cephalosporins. Therefore, isolates that are initially susceptible may become resistant within 3 to 4 days after initiation of therapy. Testing of repeat isolates may be warranted.” Some medical facilities may use this as a statement if they choose to report broad-spectrum cephalosporin susceptibilities for Serratia species. At my medical facility, we do not report broad-spectrum cephalosporin susceptibility test results for Serratia species, although S. marcescens isolates from 2008 to 2010 were 100% sensitive to ceftazidime and 97% sensitive to ceftriaxone (Table 4). An outbreak of a multiply antibiotic-resistant S. marcescens clone occurred in Italy from 2001 to 2002 and may have been due to ampC derepression or induction. The outbreak occurred among 13 patients, and 12 of the patients had been treated with various ␤-lactams before isolation of S. marcescens. The S. marcescens clone in this cluster was resistant to penicillins, aztreonam, and expanded- and broad-spectrum cephalosporins and was sensitive to carbapenems and cefepime (26). An outbreak due to S. marcescens expressing an AmpC-like

SERRATIA INFECTIONS

777

␤-lactamase, S4, was described in Taiwan from 1999 to 2003. A total of 58 strains carried this S4 ␤-lactamase, and all were recovered from patients with bloodstream infections. Strains expressing S4 were resistant to cefotaxime but not ceftazidime (420). Data on chromosomal ampC genes of other Serratia species are more limited. In one study, ␤-lactam sensitivity patterns indicated that isolates of S. liquefaciens, S. grimesii, and S. proteamaculans harbored chromosomal ampC genes (368). The sequence of the S. proteamaculans strain 568 genome indicates the presence of a chromosomal ampC genes and several other ␤-lactamases. In another study, S. ficaria, S. fonticola, S. odorifera, S. plymuthica, and S. rubidaea were shown to have chromosomal ampC genes (367). The ampC genes of S. ficaria, S. fonticola, and S. odorifera were inducible in this study, while the ampC genes of S. rubidaea and four of five strains of S. plymuthica were not (367). In a study from India of isolates recovered from different types of clinical specimens from 2007 to 2008, 25.6% of Serratia isolates produced AmpCs, 40% of which were inducible, and 60% of these isolates were derepressed mutants. Besides S. marcescens, the authors did not identify the isolates to the species level (321). There are several excellent AmpC-related reviews, including those written by Jacoby (197) and Hanson and Sanders (173). Carbapenem resistance in Serratia species. Carbapenems such as imipenem and meropenem are important antibiotics, since they are often used to treat severe infections caused by Enterobacteriaceae organisms resistant to broad-spectrum cephalosporins. Carbapenem resistance is uncommon in Serratia species (367, 368). A carbapenemase, eventually called SME-1, was first found in two S. marcescens isolates in 1982 in England (417). These isolates were both resistant to imipenem and had reduced sensitivity to meropenem (417). In addition, the two isolates were fully sensitive to broad-spectrum cephalosporins (417). Since then, two other SME-type enzymes have been described: SME-2 and SME-3 (311). The SME-type carbapenemases are class A enzymes that have a serine at the active site (311). These enzymes are not ubiquitous in S. marcescens strains, and at this point, only sporadic infections with S. marcescens isolates expressing SME carbapenemases in the United States have been described (105, 312). The SME-type carbapenemases strongly hydrolyze penicillins, narrow-spectrum cephalosporins, carbapenems, and aztreonam, weakly hydrolyze broad-spectrum cephalosporins, and are inhibited by clavulanate (311). Another chromosomal class A carbapenemase, designated SFC-1, was found in an environmental strain of S. fonticola from Portugal (180). This strain was also found to harbor a metallo-␤-lactamase called Sfh-I (180). This strain was resistant to both meropenem and imipenem (180). However, neither of the enzymes has been found in other strains of S. fonticola (180, 402). Plasmid-mediated carbapenemases have also been found in S. marcescens. From 2006 to 2007, an outbreak of 21 plasmidmediated, KPC-2-expressing S. marcescens strains was reported from China. All of the isolates were of the same clone and either were resistant to carbapenems or exhibited reduced susceptibility to carbapenems (59). KPC class carbapenemases were first discovered in K. pneumoniae in 1996, and these class A ␤-lactamases strongly hydrolyze all of the ␤-lactams and are

778

MAHLEN

plasmid mediated (311). A smaller outbreak of KPC-2-expressing S. marcescens was reported from Greece in 2008 to 2009, when three strains were recovered that had reduced susceptibility to carbapenems (385). In 2009, an S. marcescens isolate that was resistant to imipenem, meropenem, ertapenem, and doripenem was recovered from the sputum of a patient with pneumonia. The S. marcescens isolate probably acquired the carbapenemase by plasmid transfer from an E. coli isolate that the patient was also infected with; the E. coli isolate had probably previously acquired the carbapenemase from a Klebsiella pneumoniae isolate, again from the same patient. The carbapenemase from each of the bacteria was a KPC-3 enzyme (352). KPC enzymes have been found in S. marcescens on other occasions; a KPC-2 enzyme was identified from an isolate from China in 2006, and a KPC-3 enzyme was identified from an isolate from New York City in 2000 (105, 426). The appearance of different KPC enzymes in S. marcescens isolates from several different geographic locations is alarming, especially since these carbapenemases mediate such high-level resistance to carbapenems and other ␤-lactams. Another plasmid-mediated carbapenemase, GES-1, was found in all strains from 15 patients in another outbreak caused by S. marcescens in a Dutch hospital from 2002 to 2003 (106). The GES carbapenemases are also class A enzymes that are plasmid mediated (402). GES-1 exhibits low-level carbapenemase activity and was initially classified as an ESBL because it hydrolyzed penicillin and broad-spectrum cephalosporins (311, 402). Plasmid-mediated class B metallo-␤-lactamases have also been identified in S. marcescens. The metallo-␤-lactamases hydrolyze carbapenems, are not inhibited by ␤-lactamase inhibitors, are inhibited by metal ion chelators, and have zinc ions at the active site (311). There are several plasmid-borne metallo␤-lactamase genes, and the first found in S. marcescens encoded an IMP-1 enzyme (288). This enzyme, produced from an S. marcescens strain with high-level resistance to several ␤-lactam antibiotics, including imipenem and meropenem, was recovered from a patient in 1991 in Japan (288). Since then, various plasmid-mediated IMP enzymes have been found in S. marcescens several times, including from a few outbreaks (182, 303). Another type of plasmid-encoded metallo-␤-lactamase, VIM, has been found in S. marcescens (422) and S. liquefaciens (271). A survey of Serratia species from clinical isolates from India in 2007 to 2008 found that 15.4% produced metallo-␤lactamases, although the type of enzyme was not determined, and besides S. marcescens, the other Serratia species were not identified (321). Lastly, an outbreak of meropenem-resistant S. marcescens in 2005 occurred in South Korea among nine different patients. None of the isolates carried a carbapenemase, and resistance to carbapenems was probably due to overproduction of the chromosomally encoded AmpC enzyme and to loss of outer membrane protein F (OmpF) (371). Excellent reviews about carbapenemases include those written by Queenan and Bush (311) and Walther-Rasmussen and Høiby (402). ESBLs in Serratia species. The broad-spectrum cephalosporins were introduced in the early 1980s and were used to treat infections by organisms with ␤-lactamases such as TEM and SHV (300). The ESBLs are plasmid-mediated enzymes that have activity against the narrow-, expanded-,

CLIN. MICROBIOL. REV.

and broad-spectrum cephalosporins, the penicillins, and aztreonam (300). There are a wide variety of ESBLs, including TEM-, SHV-, OXA-, and CTX-M-type enzymes. There are several reports of ESBL-expressing S. marcescens isolates. In some cases, ESBL-expressing S. marcescens strains have caused outbreaks (94, 196, 281, 284, 293). S. marcescens strains most commonly carry CTX-M-type ESBLs (69, 196, 218, 273, 284, 293, 295, 414, 421) but have also been found carrying SHV (218, 281, 284, 295), TEM (218, 284, 295), and a novel ESBL, BES-1 (42). The prevalence of ESBLs in S. marcescens varies. In Taiwan, 12.2% of S. marcescens strains recovered from clinical specimens over about a 6-month period from 2001 to 2002 produced ESBLs. All of the ESBLs from this study were identified as CTX-M-3, and 33% of the patients with ESBL-producing S. marcescens died (69). In another study of S. marcescens isolates recovered from several hospitals in 2005 in Taiwan, 16% showed phenotypic ESBL production (resistance to ceftazidime, ceftriaxone, or cefepime); molecular characterization of ESBLs was not conducted (199). Rates of ESBL-producing S. marcescens from South Korea range from 12.4% (72) to 30.6% (214). In a study from Thailand, 24.1% of S. marcescens isolates recovered from 2006 to 2007 were ESBL producers; the isolates carried mixtures of CTX-M-, SHV-, and TEM-type enzymes (218). A survey of S. marcescens isolates from 2006 to 2009 in Mexico revealed that 20.5% were ESBL producers, and all of the ESBLs were SHV-type enzymes (143). In India, Rizvi and others found that 33% of Serratia species recovered from various clinical specimens from 2007 to 2008 were ESBL producers; they did not determine the type of enzymes present and did not report which species of Serratia were present besides S. marcescens (321). Several studies have been conducted in Poland to examine ESBL-producing Serratia species. In a survey from two hospitals in Danzig from 1996 to 2000, 19% of S. marcescens isolates produced ESBLs (284). Most (84%) expressed CTX-M-type enzymes (284). In one alarming national report for 2003 to 2004, enteric bacteria from 13 different hospitals in Poland were studied for ESBL production. In this study, 70.8% of S. marcescens strains were ESBL producers (122). Most (80.1%) carried CTX-M-type enzymes, while the rest produced SHV-type ESBLs. Another Polish study also showed alarming results. In this survey, 77.8% of S. marcescens isolates from 2005 from a transplantation unit exhibited phenotypic ESBL production; molecular characterization of isolates was not performed. The authors found, though, that 26.3% of S. marcescens isolates recovered from patients from other wards of the same hospital expressed phenotypic ESBL production (272). An excellent ESBL review is that written by Paterson and Bonomo (300). Quinolone Resistance in Serratia Species Quinolones target DNA gyrase and topoisomerase IV (325). DNA gyrase, encoded by gyrA and gyrB, is a type II topoisomerase that is essential for DNA replication and transcription (325). In general, Serratia species are often fairly sensitive to quinolones (367, 368). At my institution, 95% of S. marcescens strains recovered from 2008 to 2010 were sensitive to cipro-

VOL. 24, 2011

floxacin, and during this time, all (100%) strains were sensitive to levofloxacin (Table 4). Sheng and others, however, found that fluoroquinolone sensitivity decreased in S. marcescens and other Gram-negative bacteria from the mid-1980s to the late 1990s in Taiwan (348). For example, 99% of S. marcescens isolates recovered from 1985 to 1986 were sensitive to ciprofloxacin, but only 80% of isolates from 1996 to 1997 were sensitive to ciprofloxacin (348). In the two studies of Serratia susceptibilities performed by Stock and others, all of the Serratia species tested were sensitive to the quinolones, although reduced sensitivities were observed with some strains of S. marcescens and S. rubidaea (367, 368). When quinolone resistance in Serratia species does occur, it can be by a variety of mechanisms, as with other Gram-negative rods, and has most often been described for S. marcescens. S. marcescens has chromosomal determinants for quinolone resistance and also may develop resistance by acquiring plasmids or by mutation. Alterations in gyrA have commonly been shown to be involved in quinolone resistance (405). In 1991, a spontaneous ciprofloxacin-resistant mutant of an intrinsically ciprofloxacinsensitive S. marcescens isolate was recovered after incubation on medium containing 0.5 ␮g/ml of ciprofloxacin (254). This spontaneous resistance was due to a gyrA mutation (254). gyrA mutations in several quinolone-resistant Enterobacteriaceae strains, including S. marcescens, were studied by Weigel and others (405). This group found that there were several single amino acid substitutions in GyrA that enabled fluoroquinolone resistance in S. marcescens (405). Kim and others also studied quinolone-resistant S. marcescens strains and found two different single amino acid substitutions in GyrA (216). Alteration of outer membrane proteins was reported as a cause of quinolone resistance (and also resistance to aminoglycosides and some ␤-lactams) in S. marcescens in the mid-1980s (334). Omp1 appears to be the primary porin that allows ciprofloxacin entry into S. marcescens, and Omp1-deficient strains had higher MICs than those for the parent strains for several antibiotics, including ciprofloxacin and ␤-lactams such as cefoxitin, ceftriaxone, cefotaxime, and moxalactam (328). Efflux pumps are a common cause of quinolone resistance, especially in Gram-negative bacteria (31). At this point, three different chromosomally mediated efflux pumps of the resistance-nodulation-cell division (RND) family have been identified in S. marcescens: SdeAB, SdeCDE, and SdeXY. The SdeCDE pump appears to be selective and provides resistance to novobiocin (31). Norfloxacin and tetracycline are substrates for SdeXY, so this pump also appears to be fairly selective (68). The primary efflux pump of S. marcescens that utilizes quinolones as substrates appears to be SdeAB, and it provides resistance to ciprofloxacin, norfloxacin, and ofloxacin (31, 224). SdeAB also acts as an active efflux pump for chloramphenicol, sodium dodecyl sulfate, ethidium bromide, and n-hexane (224). Interestingly, it was shown that exposure of S. marcescens to cetylpyridinium chloride, a quaternary ammonium disinfectant, caused mutations in SdeAB that increased resistance to norfloxacin, biocides, and several other antibiotics (255). Another efflux pump characterized from S. marcescens, SmdAB, belongs to the ATP-binding cassette (ABC) family (257). When cloned into E. coli, this pump provided elevated MICs for several antimicrobials, including the quinolones cip-

SERRATIA INFECTIONS

779

rofloxacin, norfloxacin, ofloxacin, and nalidixic acid (257). Efflux pumps have not been well characterized for other Serratia species, but several are predicted from the genome sequence of S. proteamaculans strain 568. Another mechanism of quinolone resistance in bacteria is via the plasmid-mediated qnr genes. The qnr genes, qnrA, qnrB, qnrS, qnrC, and qnrD, code for pentapeptide repeat proteins that block quinolones from acting upon their targets. The effect of these Qnr proteins is usually low-level resistance to quinolones (253). While Qnr-mediated quinolone resistance is not often high, the presence of these determinants appears to enable further selection of more resistant mutants (253). In a 2007 study from Korea, qnr genes were found in 2.4% (4/166 strains) of S. marcescens strains; one isolate had a qnrA1 gene, two had qnrB1 genes, and one had a qnrB4 gene (296). A chromosomally mediated type of qnr gene was found in an S. marcescens isolate from Spain by Velasco and others. This qnr gene, called Smaqnr, has 80% homology to qnrB1 and was responsible for decreased ciprofloxacin susceptibility. The authors identified chromosomally carried Smaqnr in 14 other S. marcescens clinical isolates, so it may be widely distributed (394). Chromosomal qnr genes have been found in many other Gram-negative and Gram-positive bacteria (325). Recently, aac(6⬘)-Ib-cr, a variant of the aminoglycosidemodifying determinant aac(6⬘)-Ib, was found to modify ciprofloxacin by acetylation and to cause low-level resistance. The aac(6⬘)-Ib-cr gene is plasmid mediated and was shown to be additive with qnrA in determining ciprofloxacin resistance (323). To date, this plasmid-mediated gene has been found in two S. marcescens clinical isolates from South Korea. Both strains also had a plasmid-mediated qnr gene; one had qnrA1, and the other had qnrB1. The isolate with the qnrA1 gene had higher MICs for both ciprofloxacin (4 ␮g/ml) and nalidixic acid (32 ␮g/ml) than the isolate with the qnrB1 gene (0.125 ␮g/ml for ciprofloxacin and 2 ␮g/ml for nalidixic acid) (217). Rodríguez-Martínez and others provide a recent, detailed review on quinolone resistance (325). Resistance to the Tetracyclines in Serratia Species In general, many Serratia species exhibit intrinsic resistance to the tetracyclines (367, 368). All S. marcescens and S. liquefaciens isolates were resistant to tetracycline in the 2003 study by Stock and others, and most strains were resistant to other tetracyclines, such as doxycycline and minocycline (368). Thus, tetracycline, doxycycline, and minocycline are generally not good choices of therapy for S. marcescens. Resistance to the tetracyclines in Serratia has so far been described as mediated by either chromosomally mediated or plasmid-mediated efflux pumps. Some of the described chromosomally mediated efflux pumps that mediate quinolone resistance may also be responsible for tetracycline resistance. Tetracycline is a substrate for the RND pump SdeXY (68). Matsuo and others showed that the ABC pump SmdAB provided increased tetracycline resistance when it was cloned into a susceptible E. coli strain (257). In addition, the RND pump SdeAB was shown to provide an increase in tetracycline resistance after S. marcescens was exposed to cetylpyridinium chloride (255). Also, a tetracyclinespecific efflux pump, encoded by tetA(41), was identified in an

780

MAHLEN

S. marcescens strain recovered from a heavy metal-contaminated stream. The tetA(41) gene was not found on a plasmid, so it is probably located on the S. marcescens chromosome (380). Plasmid-mediated tetracycline resistance determinants have been identified in S. marcescens as well. The tetA, tetB, tetC, and tetE genes have all been found in S. marcescens strains. These genes all code for efflux pumps. Tetracycline and minocycline are substrates for TetB, but the other pumps primarily transport tetracycline (73). Tigecycline, a glycylcycline, was approved for human use in the United States in the mid-2000s. Tigecycline has shown promise against Gram-negative bacteria because it is more stable in the presence of tetracycline-specific efflux pumps such as TetA and TetB than other tetracyclines. Fritsche and others determined tigecycline susceptibilities of tetracycline-resistant Enterobacteriaceae organisms recovered from around the world from 2000 to 2004. Most of the enteric isolates were sensitive to tigecycline; however, a small percentage of S. marcescens isolates (2.4%) were resistant (138). In 2004, the Tigecycline Evaluation and Surveillance Trial (TEST) was initiated as a global survey to evaluate the effectiveness of tigecycline against Gram-negative and Gram-positive bacteria. In the United States, 96.6% of S. marcescens isolates (n ⫽ 678) in 2005 were sensitive to tigecycline; in 2006, 96.8% (n ⫽ 593) were sensitive, and in 2007, 95.8% (n ⫽ 427) were sensitive (114). The resistance of some strains of S. marcescens to tigecycline is probably due to intrinsic efflux; Hornsey and others demonstrated that upregulation of the RND efflux pump SdeXY mediates tigecycline, ciprofloxacin, and cefpirome resistance (188). More clinical data need to be collected regarding the use of tigecycline for treatment of Serratia infections. Trimethoprim-Sulfamethoxazole Resistance in Serratia Species Trimethoprim and sulfamethoxazole were first used in combination in 1968, and together they act synergistically to inhibit folic acid synthesis in bacteria. Sulfamethoxazole inhibits dihydropteroate synthetase (DHPS), an enzyme that catalyzes the formation of dihydrofolate from para-aminobenzoic acid. Trimethoprim acts on the next step of the pathway, by inhibiting the enzyme dihydrofolate reductase (DHFR); this enzyme catalyzes the conversion of dihydrofolate into tetrahydrofolate (192). Serratia species are generally thought to be susceptible to trimethoprim-sulfamethoxazole (367, 368). At my institution, all 110 S. marcescens strains recovered from clinical samples from 2008 to 2010 were sensitive to trimethoprim-sulfamethoxazole (Table 4). There are several potential mechanisms of resistance to trimethoprim and sulfamethoxazole, including cell impermeability and/or efflux pumps, intrinsically insensitive DHPS or DHFR, acquired insensitive DHPS or DHFR, and mutations, recombination events, or regulatory changes that occur in DHPS or DHFR. At least 20 transferable dhfr genes that mediate trimethoprim resistance have been described; dhfrI and different types of dhfrII are most common, especially among the Enterobacteriaceae. At this point, two transferable genes, sulI and sulII, have been found that mediate resistance to sulfonamides (192).

CLIN. MICROBIOL. REV.

While Serratia species are usually considered to be sensitive to trimethoprim-sulfamethoxazole, this may depend on the geographic area the organisms are recovered from; high resistance rates have been described over the years in several studies. In a study from Beirut, Lebanon, from 1994, Araj and others reported that 56% of Serratia species recovered from a variety of clinical sites were resistant to trimethoprim-sulfamethoxazole, compared to 12 to 48% resistance in Saudi Arabia, 50% resistance in Kuwait, and no resistance in the United States (13). From 1997 to 1999, S. marcescens isolates recovered from respiratory sites were 64 to 75% sensitive to trimethoprim-sulfamethoxazole in Italy (134). National antimicrobial resistance surveillance in Taiwan from the year 2000 indicated that 62% of S. marcescens isolates were resistant to trimethoprim-sulfamethoxazole (232). In a recent survey from Nicaragua, 27.3% of S. marcescens isolates recovered in 2008 were resistant to trimethoprim-sulfamethoxazole (45). In contrast, most (98.1%) Serratia species recovered in Canada from 2000 to 2005 were sensitive to trimethoprim-sulfamethoxazole (233). Few studies have determined the actual mechanism of resistance to trimethoprim-sulfamethoxazole in Serratia species. One study of trimethoprim-resistant Enterobacteriaceae from Greece found two S. marcescens isolates with plasmid-mediated dhfrII genes, and nine total S. marcescens isolates that were resistant to trimethoprim were recovered from urine specimens (384). Huovinen described trimethoprim resistance due to impaired permeability with S. marcescens (192). Given the facts that several efflux pumps have been identified in S. marcescens and that several more appear likely to be identified from the genome sequence of strain DB11, it is possible that trimethoprim and/or sulfonamide resistance may also be mediated by an efflux mechanism. Treatment of Serratia Species Infections Because Serratia species are intrinsically resistant to a large number of antibiotics, there are fewer treatment options for these organisms than for many other bacteria. Multiresistant Serratia strains are routinely isolated from human clinical infections, and highly resistant strains have been causative agents in many outbreaks, but a glance at Table 4 reveals that the majority of strains from different locations in the United States and Europe are sensitive to commonly reported antibiotics. Health care providers may empirically treat suspected Serratia infections with piperacillin-tazobactam, a fluoroquinolone, an aminoglycoside, and/or a carbapenem and then modify treatment based on actual susceptibility test results when available. At my institution, therapy with piperacillin-tazobactam, an aminoglycoside, and/or a carbapenem is usually successful in treating serious Serratia infections. LABORATORY IDENTIFICATION OF SERRATIA SPECIES The members of the genus Serratia are in the family Enterobacteriaceae and are not particularly difficult to cultivate from clinical specimens. Identification of Serratia species, though, can be difficult, and since species besides S. marcescens are capable of causing human infections, it is important to reliably

VOL. 24, 2011

SERRATIA INFECTIONS

identify these organisms to the species level, especially since strains of Serratia species are often multiply antibiotic resistant. As members of the Enterobacteriaceae, all Serratia species are Gram-negative rods that typically ferment glucose and are oxidase negative. Serratia species also usually reduce nitrate to nitrite and are Voges-Proskauer positive (128, 159). The mol% G⫹C content of DNA for Serratia species ranges from 52 to 60%, although the range in S. fonticola is 49 to 52% (159). In general, phenotypic systems such as the API 20E strip (bioMe´rieux), the Vitek 2 assay (bioMe´rieux), the Microscan Walk-Away test (Dade-Behring, Siemens), and the BD Phoenix test (BD Diagnostics, Sparks, MD) accurately identify several Serratia species, especially the most common species recovered from clinical specimens, S. marcescens and S. liquefaciens. Members of the genus Serratia, save S. fonticola, can generally be differentiated from most members of Klebsiella and Enterobacter on the basis of gelatin and DNase activity (128, 159). There is, however, variation among the species of the genus Serratia, and phenotypic characteristics may have to be analyzed carefully, especially if a low-percentage identification is obtained with a system or kit. Phenotypic Identification Cultural and microscopic characteristics. Serratia species generally grow well on standard clinical laboratory media, such as tryptic soy agar with 5% sheep blood (SBA), chocolate agar, and MacConkey agar (MAC). Like those of other Enterobacteriaceae, colonies will typically be fairly large after overnight incubation, i.e., about 2 mm (159). These bacteria typically grow well at 30 to 37°C, although S. plymuthica may not grow well at 37°C (159, 160). Many Serratia species are able to grow at 4 to 5°C, but not S. marcescens, S. rubidaea, and S. ureilytica (36, 159). Nonpigmented strains will often be whitish to grayish, like many other Enterobacteriaceae, and tend to be round with entire margins. The red pigmentation exhibited by many strains of S. marcescens, S. plymuthica, and S. rubidaea is probably the most striking feature of colonies of these organisms (Fig. 1). This red pigment, prodigiosin, is produced by many strains; however, nonpigmented clinical strains are commonly recovered from human specimens, and red-pigmented strains now tend to be environmental isolates (159). S. nematodiphila also produces red pigment, but it has not yet been isolated from human specimens (425). The other Serratia species do not produce prodigiosin. Prodigiosin, a prodiginine molecule, is a tripyrrole bioactive secondary metabolite that is bound to the cell envelope (32, 159). There are four main classes of prodiginines, and S. marcescens, S. plymuthica, and S. rubidaea produce a group 1 molecule, prodigiosin (410). Other organisms besides Serratia species produce prodigiosin, including other Gram-negative bacteria, such as Pseudomonas magnesiorubra and Vibrio psychroerythreus (410). The physiological role of prodigiosin is unclear, but the molecule has antibacterial and antifungal properties (32). Interestingly, prodigiosin and other prodiginines are currently being investigated as immunosuppressive agents and anticancer agents, as these compounds have immunomodulatory capabilities (410). As a secondary metabolite, prodigiosin is produced in the later stages of growth and is

781

sensitive to several environmental factors, including temperature, pH, the availability of light, the amount of oxygen present, and the availability of inorganic ions, various amino acids, and carbohydrates (32, 159, 410). Thus, the pigment may not be produced by particular strains unless certain environmental conditions are met. Prodigiosin-producing strains of S. marcescens belong to biogroups A1, A2, and A6 (159). S. marcescens strains that belong to nonpigmented biogroups such as A5 and A8 are frequently isolated from human specimens; ubiquitous, environmental strains often belong to biogroups A3 and A4 (159). Nonpigmented strains recovered from human clinical specimens started appearing in the late 1950s and were readily recognized by the early to mid-1960s (81, 127, 161, 231). The pigment may be red, pink, magenta, or orange, depending on cultural conditions, and colonies may be pigmented entirely or partially (159). Figure 1 shows the same S. marcescens strain inoculated onto MAC (Fig. 1A), trypticase soy yeast agar (TSY) (Fig. 1B), and sheep blood agar (Fig. 1C). The colonies of this particular isolate on MAC are bright red, and they are more orange on TSY. It is important that bench technologists do not confuse the red-pigmented colonies on MAC with lactose fermentation. Prodigiosin is a nonsoluble pigment, so it does not diffuse in agar. When cultures of S. marcescens, S. plymuthica, and S. rubidaea are incubated at 37°C, pigmentation may not appear, but it may form at 30°C (160). Occasional strains of S. marcescens biogroup A4 produce another pigment, pyrimine (159). Pyrimine has been described as both pink (159) and reddish violet (195). Pyrimine contains ferrous iron and may have properties that mimic those of superoxide dismutase (195). Some Serratia species also produce particular odors when cultured on solid media. A potato-like odor is produced by S. ficaria, S. odorifera, and some strains of S. rubidaea (141, 165, 167). The potato-like odor is due to pyrazines produced by these species (141). In addition, all of the other Serratia species are sometimes described as having a fishy-urinary odor due to trimethylamine and/or ammonia production (159). Cells of Serratia are microscopically rod-like with rounded ends and range from 0.9 to 2.0 ␮m in length and from 0.5 to 0.8 ␮m in width (159). Like some other members of the Enterobacteriaceae, they may have a bipolar, or “safety pin,” appearance on Gram staining, where the ends of the cells stain darker than the middle. Most strains of all Serratia species are motile, typically with peritrichous flagella (159), although S. nematodiphila has a single polar flagellum (425). Identification of S. marcescens. S. marcescens, the species most likely to be recovered from clinical specimens, is well known as one of the few members of the Enterobacteriaceae that produces DNase, lipase, and gelatinase (128, 159). S. marcescens does not usually ferment lactose, although pigmented strains may initially appear to be lactose fermenters on MAC without a precipitate around colonies (Fig. 1A). S. marcescens does not produce indole, is lysine and ornithine decarboxylase positive, and is arginine dihydrolase negative. In addition, S. marcescens ferments sucrose and D-sorbitol but does not ferment L-arabinose or raffinose. S. marcescens can be differentiated from pigmented strains of both S. rubidaea and S. plymuthica by ornithine decarboxylase activity and a lack of L-arabinose and raffinose fermentation. There are several S.

ND ND ⫹ ND ⫺ ⫺ ⫺ ⫹ ⫹ ⫹ ⫹ ⫺ ND ⫺ ⫹ ⫹ ⫹ ⫹ ⫹ ND ⫹ ⫺ ⫺ ⫺ ⫹ ⫹ ⫹ ⫹ ND ⫹ ⫹ ⫹ ND ⫺ ND ND ⫺ ⫺ ⫺ ⫹ ⫺ ⫹ ⫹ ⫹ ND ND ⫹ ⫺ b

a

Characteristics were compiled from several references (36, 128, 146, 159, 425). ND, not determined; V, variable reaction. S. odorifera biotype 1 is ornithine decarboxylase positive and ferments sucrose, while biotype 2 is ornithine decarboxylase negative and does not ferment sucrose.

⫹ ⫹ ⫹ ⫹ ⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹ ⫹ ⫹ ⫹ ⫹ V ⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹ ⫺ ⫹ V ⫹ ⫹ ⫹ ⫺ ⫹ ⫺ ⫹ V ⫺ ⫺ ⫹ Vb ⫹ ⫺ ⫹ ⫹ Vb ⫹ ⫹ ⫹ ⫹ V V ⫺ ⫺ ⫺ V ⫺ ⫹ ⫺ ⫹ ⫺ ⫹ ⫺ ⫺ ⫹ ⫹ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹ ⫹ ⫹ ⫹ ⫹ ⫹ V ⫹ ⫹ ⫹ ⫹ ⫺ ⫹ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹ ⫺ V ⫹ ⫹ ⫹ ⫹ ⫹ ⫹ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹ ⫹ ⫹ ⫺ V V ⫹ ⫹ ⫹ ⫹ ⫹ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹ ⫹ ⫹ ⫺ V V ⫹ ⫹ ⫹ ⫹ ⫹ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹ ⫹ ⫹ ⫺ V ⫹ ⫹ ⫹ ⫹ ⫹ ⫹ V ⫺ ⫺ ⫺ ⫺ ⫹ ⫹ ⫺ ⫺ ⫺ ⫹ ⫹ DNase Gelatinase Lipase (Tween 80 hydrolysis) Lipase (corn oil hydrolysis) Prodigiosin production Potato odor Indole Urease Arginine dihydrolase Lysine decarboxylase Ornithine decarboxylase L-Arabinose fermentation D-Dulcitol fermentation Lactose fermentation D-Sorbitol fermentation Sucrose fermentation

⫹ ⫹ ⫹ ⫹ ⫺ ⫺ ⫺ ⫺ ⫹ ⫹ ⫹ ⫹ ⫺ ⫹ ⫹ ⫹

S. nematodiphila S. glossinae S. entomophila S. plymuthica S. odorifera S. rubidaea S. fonticola S. ficaria S. quinivorans S. proteamaculans S. grimesii S. liquefaciens S. marcescens Characteristic

marcescens biogroups and biovars; their differential characteristics are summarized in the current edition of Bergey’s Manual of Systematic Bacteriology (159). See Table 5 for a selected list of characteristics useful for identifying S. marcescens and other Serratia isolates to the species level. Identification of Serratia species. In addition to S. marcescens, most strains of species of the genus Serratia are positive for DNase production and gelatin hydrolysis (128, 159). S. fonticola is negative for these tests, though, is Voges-Proskauer negative, and is phenotypically much different from other Serratia species (145). Except for many strains of S. odorifera, Serratia species do not usually produce indole (128, 159), and only S. ureilytica and S. glossinae, both of which have not been implicated in human infections, produce urease (36, 146). Most strains of all species utilize citrate, hydrolyze esculin, hydrolyze corn oil (lipase), and are H2S negative (128, 159, 425). S. odorifera is the only species that does not hydrolyze Tween 80 (159). There are also general patterns of carbon source utilization for the genus. Most strains of each species utilize maltose, D-mannitol, D-mannose, and trehalose, while dulcitol is not utilized by any species except for S. fonticola (128, 159). There are biotypes of S. entomophila, S. grimesii, S. liquefaciens, S. odorifera, S. proteamaculans, S. quinivorans, and S. rubidaea, and differential characteristics for these biotypes are listed in the current edition of Bergey’s Manual of Systematic Bacteriology (159). See Table 5 for selected phenotypic characteristics for each Serratia species; for more complete characteristics, consult the current editions of Bergey’s Manual of Systematic Bacteriology (159) and the Manual of Clinical Microbiology (128) and the papers with descriptions of S. ureilytica (36), S. glossinae (146), and S. nematodiphila (425). A brief summary of key characteristics of Serratia species (except for S. marcescens) follows. (i) S. liquefaciens. S. liquefaciens isolates are not pigmented and produce DNase, gelatinase, and lipase. Most strains are lysine decarboxylase and ornithine decarboxylase positive. S. liquefaciens strains are indole, urease, and arginine dihydrolase negative. This organism is part of the S. liquefaciens complex, along with S. grimesii, S. proteamaculans, and S. quinivorans. (ii) S. grimesii. S. grimesii is part of the S. liquefaciens complex and is not pigmented. Isolates produce DNase, gelatinase, and lipase and are arginine dihydrolase, lysine decarboxylase, and ornithine decarboxylase positive. S. grimesii ferments lactose. (iii) S. proteamaculans. S. proteamaculans is in the S. liquefaciens complex. It is not pigmented and produces DNase, gelatinase, and lipase. It is lysine decarboxylase and ornithine decarboxylase positive. S. proteamaculans is indole, urease, and arginine dihydrolase negative. (iv) S. quinivorans. Like S. liquefaciens, S. grimesii, and S. proteamaculans, S. quinivorans is in the S. liquefaciens complex. S. quinivorans produces DNase, gelatinase, and lipase. It is not pigmented and is indole, urease, and arginine dihydrolase negative. S. quinivorans is lysine decarboxylase and ornithine decarboxylase positive. (v) S. ficaria. S. ficaria colonies are nonpigmented and produce a potato-like odor. This organism produces DNase, gelatinase, and lipase. S. ficaria isolates are indole, urease, arginine dihydrolase, lysine decarboxylase, and ornithine decarboxylase negative.

CLIN. MICROBIOL. REV. S. ureilytica

MAHLEN

TABLE 5. Phenotypic characteristics of members of the genus Serratiaa

782

VOL. 24, 2011

SERRATIA INFECTIONS

(vi) S. fonticola. S. fonticola differs from the other species in the genus because most strains ferment D-dulcitol and do not produce DNase and gelatinase. S. fonticola produces lipase, is not pigmented, and is indole and urease negative. This organism is lysine decarboxylase and ornithine decarboxylase positive, usually ferments lactose, and is arginine dihydrolase negative. (vii) S. rubidaea. S. rubidaea may be pigmented, and some strains also have a potato-like odor. It produces DNase, gelatinase, and lipase and ferments lactose. S. rubidaea may be lysine decarboxylase positive but is arginine dihydrolase and orthinine decarboxylase negative. This organism is indole and urease negative and does not ferment D-sorbitol. (viii) S. odorifera. S. odorifera does not produce pigment and has a potato-like odor. It produces DNase and gelatinase, but it is the only Serratia species that does not hydrolyze Tween 80. Some strains of S. odorifera are indole positive. This organism is urease negative and lysine decarboxylase positive and usually ferments lactose. There are two biotypes, and biotype 1 is ornithine decarboxylase positive and ferments sucrose; biotype 2 is ornithine decarboxylase negative and does not ferment sucrose. (ix) S. plymuthica. Like S. marcescens and S. rubidaea, S. plymuthica may be pigmented. It produces DNase, gelatinase, and lipase and ferments lactose. S. plymuthica is indole, urease, arginine dihydrolase, lysine decarboxylase, and ornithine decarboxylase negative. (x) S. entomophila. S. entomophila is not pigmented. S. entomophila produces DNase, gelatinase, and lipase but is indole, urease, arginine dihydrolase, lysine decarboxylase, and ornithine decarboxylase negative. In addition, it does not ferment L-arabinose, lactose, or D-sorbitol. (xi) S. glossinae. Laboratory identification data on S. glossinae are limited, since characteristics for only one strain have been determined, and not all tests were performed (146). S. glossinae is not pigmented and is the only Serratia species that does not produce gelatinase; DNase and lipase production were not determined for this isolate. S. glossinae is urease positive and indole and arginine dihydrolase negative. This organism is lysine decarboxylase and ornithine decarboxylase positive. S. glossinae is the only Serratia species besides S. odorifera biotype 2 that does not ferment sucrose. (xii) S. nematodiphila. As with S. glossinae, only one isolate of S. nematodiphila has been characterized (425). S. nematodiphila is red pigmented and is also fluorescent. It produces DNase, gelatinase, and lipase and is arginine dihydrolase, lysine decarboxylase, and ornithine decarboxylase positive. S. nematodiphila ferments lactose and is indole and urease negative. (xiii) S. ureilytica. Only one isolate of S. ureilytica has been characterized, similar to both S. glossinae and S. nematodiphila (36). S. ureilytica produces lipase, but DNase and gelatinase were not tested. It is the only Serratia species besides S. glossinae that produces urease. This organism is arginine dihydrolase, lysine decarboxylase, and ornithine decarboxylase positive. S. ureilytica is indole negative, and like S. marcescens and S. entomophila, it does not ferment L-arabinose. Molecular Identification 16S rRNA gene sequencing is a method that efficiently distinguishes Serratia species. Figure 2 shows a dendrogram of the 16S rRNA gene sequences of the current species in the genus

783

Serratia, constructed by the neighbor-joining method in MicroSeq software (Applied Biosystems). 16S rRNA gene sequences of type strains were obtained from GenBank and are up to date through July 2011. The GenBank accession numbers used in the construction of the dendrogram are listed next to the species in Fig. 2. Links to each Serratia species type strain 16S rRNA gene sequence are available at the List of Prokaryotic Names with Standing in Nomenclature website (http://www .bacterio.cict.fr/s/serratia.html). 16S rRNA gene sequencing will differentiate Serratia species, including the members of the S. liquefaciens group (Fig. 2). A comparison of the sequences used in the construction of the dendrogram in Fig. 2 reveals that S. liquefaciens and S. grimesii differ by 6 bases, and S. proteamaculans and S. quinivorans also differ by 6 bases. In both cases, 16S rRNA gene sequencing would provide enough discrimination to identify these organisms. 16S rRNA gene sequencing, however, probably does not differentiate between biotypes or biogroups of Serratia species, including those of S. entomophila, S. grimesii, S. liquefaciens, S. marcescens, S. odorifera, S. proteamaculans, S. quinivorans, and S. rubidaea. The definition of what level of sequencing discrimination defines a species has not been determined, although a 0.5% to 1% difference is often used (80). In order to identify Serratia species biotypes, the differential characteristics listed in the current edition of Bergey’s Manual of Systematic Bacteriology may be used (159). Since the more common species in the genus are typically identified well with phenotypic systems, 16S rRNA gene sequencing does not have to be used often in clinical laboratories to determine the identity of problematic organisms. However, if a low-percentage identity is obtained with a system, 16S rRNA gene sequencing is useful for identification of the Serratia species. ACKNOWLEDGMENTS I thank the clinical microbiology staff of the Department of Pathology at Madigan Healthcare System, in particular Jessica Cromheecke and Paul Mann, for their assistance. The views expressed in this paper are those of the author and do not reflect the official policy or position of the Department of the Army, Department of Defense, or U.S. Government. REFERENCES 1. Aïtoff, M., M. Dion, and H. Dobkevitch. 1936. Bacillus prodigiosus pathoge`ne pour les animoux: endotoxine et reaction de Shwartzman. C. R. Soc. Biol. 123:375–376. (In French.) 2. Ajithkumar, B., V. P. Ajithkumar, R. Iriye, Y. Doi, and T. Sakai. 2003. Spore-forming Serratia marcescens subsp. sakuensis subsp. nov., isolated from a domestic wastewater treatment tank. Int. J. Syst. Evol. Microbiol. 53:253–258. 3. Alexandrakis, G., E. C. Alfonso, and D. Miller. 2000. Shifting trends in bacterial keratitis in south Florida and emerging resistance to fluoroquinolones. Ophthalmology 107:1497–1502. 4. Al Jarousha, A. M., I. A. El Qouqa, A. H. N. El Jadba, and A. S. Al Afifi. 2008. An outbreak of Serratia marcescens septicaemia in neonatal intensive care unit in Gaza City, Palestine. J. Hosp. Infect. 70:119–126. 5. Allison, C., N. Coleman, P. L. Jones, and C. Hughes. 1992. Ability of Proteus mirabilis to invade human urothelial cells is coupled to motility and swarming differentiation. Infect. Immun. 60:4740–4746. 6. Amaya, et al. 2010. Antibiotic resistance patterns in gram-negative and gram-positive bacteria causing septicemia in newborns in Leo ´n, Nicaragua: correlation with environmental samples. J. Chemother. 22:25–29. 7. Anagnostakis, D., J. Fitsialos, C. Koutsia, J. Messaritakis, and N. Matsaniotis. 1981. A nursery outbreak of Serratia marcescens infection. Evidence of a single source of contamination. Am. J. Dis. Child. 135:413–414. 8. Anahory, T., H. Darbas, O. Ongaro, H. Jean-Pierre, and P. Mion. 1998. Serratia ficaria: a misidentified or unidentified rare cause of human infections in fig tree culture zones. J. Clin. Microbiol. 36:3266–3272.

784

MAHLEN

9. Anonymous. 1906. Report on an investigation of the ventilation of the debating chamber of the House of Commons. Appendix to parliamentary paper Cd. 3035. HMSO, London, England. 10. Anonymous. 1969. Serratia septicaemia. Br. Med. J. 4:756–757. 11. Anonymous. 1977. Hearings before the Subcommittee on Health and Scientific Research of the Committee on Human Resources. Biological testing involving human subjects by the Department of Defense, 1977. U.S. Government Printing Office, Washington, DC. 12. Anonymous. 2010. Annual epidemiological report on communicable diseases in Europe 2010. European Centre for Disease Prevention and Control, Stockholm, Sweden. 13. Araj, G. F., M. M. Uwaydah, and S. Y. Alami. 1994. Antimicrobial susceptibility patterns of bacterial isolates at the American University Medical Center in Lebanon. Diagn. Microbiol. Infect. Dis. 20:151–158. 14. Archibald, L. K., et al. 1997. Serratia marcescens outbreak associated with extrinsic contamination of 1% chlorxylenol soap. Infect. Control Hosp. Epidemiol. 18:704–709. 15. Ariel, I., I. Arad, and D. Soffer. 1986. Autopsy findings of Serratia meningoencephalitis in infants. Pediatr. Pathol. 6:351–358. 16. Aronson, J. D., and I. Alderman. 1943. The occurrence and bacteriological characteristics of S. marcescens from a case of meningitis. J. Bacteriol. 46:261–267. 17. Arroyo, J. C., et al. 1981. Clinical, epidemiologic and microbiologic features of a persistent outbreak of amikacin-resistant Serratia marcescens. Infect. Control 2:367–372. 18. Arslan, U., et al. 2010. Serratia marcescens sepsis outbreak in a neonatal intensive care unit. Pediatr. Int. 52:208–212. 19. Arzese, A., G. A. Botta, G. P. Gesu, and G. Schito. 1988. Evaluation of a computer-assisted method of analysing SDS-PAGE protein profiles in tracing a hospital outbreak of Serratia marcescens. J. Infect. 17:35–42. 20. Ashelford, K. E., J. C. Fry, M. J. Bailey, and M. J. Day. 2002. Characterization of Serratia isolates from soil, ecological implications and transfer of Serratia proteamaculans subsp. quinovora Grimont et al. 1983 to Serratia quinivorans corrig., sp. nov. Int. J. Syst. Evol. Microbiol. 52:2281–2289. 21. Assadian, O., et al. 2002. Nosocomial outbreak of Serratia marcescens in a neonatal intensive care unit. Infect. Control Hosp. Epidemiol. 23:457–461. 22. Atlee, W. E., R. P. Burns, and M. Oden. 1970. Serratia marcescens keratoconjunctivitis. Am. J. Ophthalmol. 70:31–33. 23. Aucken, H. M., and T. L. Pitt. 1998. Different O and K serotype distributions among clinical and environmental strains of Serratia marcescens. J. Med. Microbiol. 47:1097–1104. 24. Aucken, H. M., and T. L. Pitt. 1998. Antibiotic resistance and putative virulence factors of Serratia marcescens with respect to O and K serotypes. J. Med. Microbiol. 47:1105–1113. 25. Badenoch, P. R., A. L. Thom, and D. J. Coster. 2002. Serratia ficaria endophthalmitis. J. Clin. Microbiol. 40:1563–1564. 26. Bagattini, M., et al. 2004. A nosocomial outbreak of Serratia marcescens producing inducible AmpC-type beta-lactamase enzyme and carrying antimicrobial resistance genes within a class 1 integron. J. Hosp. Infect. 56: 29–36. 27. Barnum, D. A., E. L. Thackeray, and N. A. Fish. 1958. An outbreak of mastitis caused by Serratia marcescens. Can. J. Comp. Med. Vet. Sci. 22: 392–395. 28. Bates, C. J., and R. Pearse. 2005. Use of hydrogen peroxide vapour for environmental control during a Serratia outbreak in a neonatal intensive care unit. J. Hosp. Infect. 61:364–366. 29. Baya, A. M., A. E. Toranzo, B. Lupiani, Y. Santos, and F. M. Hetrick. 1992. Serratia marcescens: a potential pathogen for fish. J. Fish Dis. 15:15–26. 30. Beck-Sague, C. M., and W. R. Jarvis. 1989. Epidemic bloodstream infections associated with pressure transducers: a persistent problem. Infect. Control Hosp. Epidemiol. 10:54–59. 31. Begic, S., and E. A. Worobec. 2008. The role of the Serratia marcescens SdeAB multidrug efflux pump and TolC homologue in fluoroquinolone resistance studied via gene-knockout mutagenesis. Microbiology 154:454– 461. 32. Bennett, J. W., and R. Bentley. 2000. Seeing red: the story of prodigiosin. Adv. Appl. Microbiol. 47:1–32. 33. Bennett, S. N., et al. 1995. Postoperative infections traced to contamination of an intravenous anesthetic, propofol. N. Engl. J. Med. 333:147–154. 34. Bernard, L. A., and W. C. Sutton. 1960. Infection due to chromobacteria. Report of a case of pneumonia due to Chromobacterium prodigiosum successfully treated with kanamycin. Arch. Intern. Med. 105:311–315. 35. Berthelot, P., et al. 1999. Investigation of a nosocomial outbreak due to Serratia marcescens in a maternity hospital. Infect. Control Hosp. Epidemiol. 20:233–236. 36. Bhadra, B., P. Roy, and R. Chakraborty. 2005. Serratia ureilytica, sp. nov., a novel urea-utilizing species. Int. J. Syst. Evol. Microbiol. 55:2155–2158. 37. Bizio, B. 1823. Lettera di Bartolomeo Bizio al chiarissimo canonico Angelo Bellani sopra il fenomeno della polenta porporina. Bibl. Ital. G. Lett. Sci. Art. (Anno VIII) 30:275–295. (In Italian.) 38. Blossom, D., et al. 2009. Multistate outbreak of Serratia marcescens blood-

CLIN. MICROBIOL. REV.

39. 40. 41.

42.

43. 44.

45.

46.

47.

48.

49. 50.

51. 52.

53.

54.

55.

56.

57.

58. 59.

60. 61.

62. 63.

64. 65.

66. 67.

stream infections caused by contamination of prefilled heparin and isotonic sodium chloride solution syringes. Arch. Intern. Med. 169:1705–1711. Bollet, C., M. Gainnier, J.-M. Sainty, P. Orhesser, and P. de Mico. 1991. Serratia fonticola isolated from a leg abscess. J. Clin. Microbiol. 29:834–835. Bollet, C., et al. 1993. Fatal pneumonia due to Serratia proteamaculans subsp. quinovora. J. Clin. Microbiol. 31:444–445. Bollmann, R., et al. 1989. Nosocomial infections due to Serratia marcescens—clinical findings, antibiotic susceptibility patterns and fine typing. Infection 17:294–300. Bonnet, R., et al. 2000. A novel class A extended-spectrum ␤-lactamase (BES-1) in Serratia marcescens isolated in Brazil. Antimicrob. Agents Chemother. 44:3061–3068. Bosi, C., et al. 1996. Serratia marcescens nosocomial outbreak due to contamination of hexetidine solution. J. Hosp. Infect. 33:217–224. Boulton, F. E., S. T. Chapman, and T. H. Walsh. 1998. Fatal reaction to transfusion of red-cell concentrate contaminated with Serratia liquefaciens. Transfus. Med. 8:15–18. Bours, P. H. A., et al. 2010. Increasing resistance in community-acquired urinary tract infections in Latin America, five years after the implementation of national therapeutic guidelines. Int. J. Infect. Dis. 14:e770–e774. Bozhokina, E., S. Khaitlina, and T. Adam. 2008. Grimelysin, a novel metalloprotease from Serratia grimesii, is similar to ECP32. Biochem. Biophys. Res. Commun. 367:888–892. Bozhokina, E. S., et al. 2010. Bacterial invasion of eukaryotic cells can be mediated by actin-hydrolysing metalloproteases grimelysin and protealysin. Cell Biol. Int. 35:111–118. Braver, D. J., G. J. Hauser, L. Berns, Y. Siegman-Igra, and B. Muhlbauer. 1987. Control of a Serratia marcescens outbreak in a maternity hospital. J. Hosp. Infect. 10:129–137. Breed, R. S., and M. E. Breed. 1924. The type species of the genus Serratia, commonly known as Bacillus prodigiosus. J. Bacteriol. 9:545–557. Brooks, H. J., T. J. Chambers, and S. Tabaqchali. 1979. The increasing isolation of Serratia species from clinical specimens. J. Hyg. (London) 82:31–40. Brouillard, J. A., W. Hansen, and A. Compere. 1984. Isolation of Serratia ficaria from human clinical specimens. J. Clin. Microbiol. 19:902–904. Buffet-Bataillon, S., et al. 2009. Outbreak of Serratia marcescens in a neonatal intensive care unit: contaminated unmedicated liquid soap and risk factors. J. Hosp. Infect. 72:17–22. Bullock, D. W., et al. 1982. Outbreaks of hospital infection in southwest England caused by gentamicin-resistant Serratia marcescens. J. Hosp. Infect. 3:263–273. Burke, G. R., B. B. Normark, C. Favret, and N. A. Moran. 2009. Evolution and diversity of facultative symbionts from the aphid subfamily Lachninae. Appl. Environ. Microbiol. 75:5328–5335. Burke, G. R., and N. A. Moran. 25 January 2011. Massive genomic decay in Serratia symbiotica, a recently evolved symbiont of aphids. Genome Biol. Evol. doi:10.1093/gbe/evr002. Burket, L. W., and C. G. Burn. 1937. Bacteremias following dental extraction: demonstration of source of bacteria by means of a non-pathogen (Serratia marcescens). J. Dent. Res. 16:521–530. Burkholder, W. H. 1948. Genus I. Erwinia Winslow et al., p. 463–478. In R. S. Breed, E. G. D. Murray, and A. P. Hitchens (ed.), Bergey’s manual of determinative bacteriology, 6th ed. The Williams and Wilkins Co., Baltimore, MD. Cabrera, H. A. 1969. An outbreak of Serratia marcescens, and its control. Arch. Intern. Med. 123:650–655. Cai, J. C., H. W. Zhou, R. Zhang, and G. X. Chen. 2008. Emergence of Serratia marcescens, Klebsiella pneumoniae, and Escherichia coli isolates possessing the plasmid-mediated carbapenem-hydrolyzing ␤-lactamase KPC-2 in intensive care units of a Chinese hospital. Antimicrob. Agents Chemother. 52:2014–2018. Campbell, J. R., T. Diacovo, and C. J. Baker. 1992. Serratia marcescens meningitis in neonates. Pediatr. Infect. Dis. J. 11:881–886. Cann, K. J., D. Johnstone, and A. Skene. 1987. An outbreak of Serratia marcescens infection following urodynamic studies. J. Hosp. Infect. 9:291– 293. Carrero, P., et al. 1995. Report of six cases of human infection by Serratia plymuthica. J. Clin. Microbiol. 33:275–276. Casolari, C., et al. 2005. A simultaneous outbreak of Serratia marcescens and Klebsiella pneumoniae in a neonatal intensive care unit. J. Hosp. Infect. 61:312–320. Ceccaldi, B., et al. 1999. Serratia odorifera biovar 1 bronchial infection. Rev. Med. Interne 20:84–85. (In French.) Champion, H. M., P. M. Bennett, D. A. Lewis, and D. S. Reeves. 1988. Cloning and characterization of an AAC(6⬘) gene from Serratia marcescens. J. Antimicrob. Chemother. 22:587–596. Chaudhuri, A. K., and C. F. Booth. 1992. Outbreak of chest infections with Serratia marcescens. J. Hosp. Infect. 22:169–170. Chemaly, R. F., D. B. Rathod, and I. I. Raad. 2009. A tertiary care cancer center experience of the 2007 outbreak of Serratia marcescens bloodstream

VOL. 24, 2011

68.

69.

70. 71. 72.

73.

74.

75.

76.

77.

78. 79. 80.

81. 82.

83.

84. 85. 86. 87. 88.

89. 90. 91.

92. 93.

94.

95.

96. 97.

infection due to prefilled syringes. Infect. Control Hosp. Epidemiol. 30: 1237–1238. Chen, J., T. Kuroda, M. N. Huda, T. Mizushima, and T. Tsuchiya. 2003. An RND-type multidrug efflux pump SdeXY from Serratia marcescens. J. Antimicrob. Chemother. 52:176–179. Cheng, K. C., Y. C. Chuang, L. T. Wu, G. C. Huang, and W. L. Yu. 2006. Clinical experiences of the infections caused by extended-spectrum ␤-lactamase-producing Serratia marcescens at a medical center in Taiwan. Jpn. J. Infect. Dis. 59:147–152. Cheng, K. H., et al. 1999. Incidence of contact-lens-associated microbial keratitis and its related morbidity. Lancet 354:181–185. Chmel, H. 1988. Serratia odorifera biogroup 1 causing an invasive human infection. J. Clin. Microbiol. 26:1244–1245. Choi, S. H., et al. 2007. Prevalence, microbiology, and clinical characteristics of extended-spectrum ␤-lactamase-producing Enterobacter spp., Serratia marcescens, Citrobacter freundii, and Morganella morganii in Korea. Eur. J. Clin. Microbiol. Infect. Dis. 26:557–561. Chopra, I., and M. Roberts. 2001. Tetracycline antibiotics: mode of action, applications, molecular biology, and epidemiology of bacterial resistance. Microbiol. Mol. Biol. Rev. 65:232–260. Christensen, G. D., et al. 1982. Epidemic Serratia marcescens in a neonatal intensive care unit: importance of the gastrointestinal tract as a reservoir. Infect. Control 3:127–133. Chuang, T.-Y., C.-P. Chuang, H.-H. Cheng, and P. R. Hsueh. 2007. Aortic valve infective endocarditis caused by Serratia liquefaciens. J. Infect. 54: e161–e163. Cimolai, N., C. Trombley, D. Wensley, and J. LeBlanc. 1997. Heterogenous Serratia marcescens genotypes from a nosocomial pediatric outbreak. Chest 111:194–197. Civen, R., et al. 2006. Outbreak of Serratia marcescens infections following injection of betamethasone compounded at a community pharmacy. Clin. Infect. Dis. 43:831–837. Clark, R. B., and J. M. Janda. 1985. Isolation of Serratia plymuthica from a human burn site. J. Clin. Microbiol. 21:656–657. Clarke, B. 1976. Infection in the intensive care unit. Aust. N. Z. J. Surg. 46:318–321. Clarridge, J. E., III. 2004. Impact of 16S rRNA gene sequence analysis for identification of bacteria on clinical microbiology and infectious diseases. Clin. Microbiol. Rev. 17:840–862. Clayton, E., and A. von Graevenitz. 1966. Nonpigmented Serratia marcescens. JAMA 197:1059–1064. CLSI. 2011. Performance standards for antimicrobial susceptibility testing; 21st informational supplement. CLSI document M100–S21. Clinical and Laboratory Standards Institute, Wayne, PA. Cohen, A. L., et al. 2008. Outbreak of Serratia marcescens bloodstream and central nervous system infections after interventional pain management procedures. Clin. J. Pain 24:374–380. Cole, L. A. 1988. Clouds of secrecy: introduction, p. 3–10. In Clouds of secrecy. Rowman & Littlefield Publishers, Inc., Lanham, MD. Cole, L. A. 1988. Edward Nevin and the spraying of San Francisco, p. 75–84. In Clouds of secrecy. Rowman & Littlefield Publishers, Inc., Lanham, MD. Cole, L. A. 1988. The trial, p. 85–104. In Clouds of secrecy. Rowman & Littlefield Publishers, Inc., Lanham, MD. Coleman, D., et al. 1984. Simultaneous outbreaks of infection due to Serratia marcescens in a general hospital. J. Hosp. Infect. 5:270–282. Cook, L. N., R. S. Davis, and B. H. Stover. 1980. Outbreak of amikacinresistant Enterobacteriaceae in an intensive care nursery. Pediatrics 65:264– 268. Cook, M. A., and J. J. Lopez, Jr. 1998. Serratia odorifera biogroup 1: an emerging pathogen. J. Am. Osteopath. Assoc. 98:505–507. Cooper, R. L., and I. J. Constable. 1977. Infective keratitis in soft contact lens wearers. Br. J. Ophthalmol. 61:250–254. Coulthurst, S. J., N. R. Williamson, A. K. Harris, D. R. Spring, and G. P. Salmond. 2006. Metabolic and regulatory engineering of Serratia marcescens: mimicking phage-mediated horizontal acquisition of antibiotic biosynthesis and quorum-sensing capacities. Microbiology 152:1899–1911. Cowan, S. 1956. “Ordnung in das chaos” Migula. Can. J. Microbiol. 2:212– 219. Craven, P. C., J. H. Jorgensen, R. L. Kaspar, and D. J. Drutz. 1977. Amikacin therapy of patients with multiply antibiotic-resistant Serratia marcescens infections: development of increasing resistance during therapy. Am. J. Med. 62:902–910. Crivaro, V., et al. 2007. Risk factors for extended-spectrum ␤-lactamaseproducing Serratia marcescens and Klebsiella pneumoniae acquisition in a neonatal intensive care unit. J. Hosp. Infect. 67:135–141. Cullen, M. M., A. Trail, M. Robinson, M. Keaney, and P. R. Chadwick. 2005. Serratia marcescens outbreak in a neonatal intensive care unit prompting review of decontamination of laryngoscopes. J. Hosp. Infect. 59:68–70. Cumming, J. G. 1920. Sputum-borne disease transmission with epidemiologic and bacteriologic research. Mil. Surg. 46:150–165. Dalamaga, M., M. Pantelaki, K. Karmaniolas, A. Matekovits, and K.

SERRATIA INFECTIONS

98. 99.

100.

101. 102.

103.

104. 105.

106.

107.

108. 109.

110.

111. 112.

113.

114.

115. 116.

117.

118. 119.

120.

121.

122.

123.

124.

785

Daskalopoulou. 2008. Cutaneous abscess and bacteremia due to Serratia ficaria. J. Eur. Acad. Dermatol. Venereol. 22:1388–1389. Darbas, H., H. Jean-Pierre, and J. Paillisson. 1994. Case report and review of septicemia due to Serratia ficaria. J. Clin. Microbiol. 32:2285–2288. Das, S., H. Sheorey, H. R. Taylor, and R. B. Vajpayee. 2007. Association between cultures of contact lens and corneal scraping in contact lens related microbial keratitis. Arch. Ophthalmol. 125:1182–1185. David, M. D., T. M. A. Weller, P. Lambert, and A. P. Fraise. 2006. An outbreak of Serratia marcescens on the neonatal unit: a tale of two clones. J. Hosp. Infect. 63:27–33. Davis, J. T., E. Foltz, and W. S. Blakemore. 1970. Serratia marcescens: a pathogen of increasing importance. JAMA 214:2190–2192. Debast, S. B., W. J. Melchers, A. Voss, J. A. Hoogkamp-Kortanje, and J. F. Meis. 1995. Epidemiological survey of an outbreak of multiresistant Serratia marcescens by PCR-fingerprinting. Infection 23:267–271. de Boer, M. G., et al. 2008. Multifactorial origin of high incidence of Serratia marcescens in a cardio-thoracic ICU: analysis of risk factors and epidemiological characteristics. J. Infect. 56:446–453. del Pozo, J. L., and R. Patel. 2007. The challenge of treating biofilmassociated bacterial infections. Clin. Pharmacol. Ther. 82:204–209. Deshpande, L. M., P. R. Rhomberg, H. S. Sader, and R. N. Jones. 2006. Emergence of serine carbapenemases (KPC and SME) among clinical strains of Enterobacteriaceae isolated in the United States Medical Centers: report from the MYSTIC Program (1999–2005). Diagn. Microbiol. Infect. Dis. 56:367–372. de Vries, J. J., et al. 2006. Outbreak of Serratia marcescens colonization and infection traced to a healthcare worker with long-term carriage on the hands. Infect. Control Hosp. Epidemiol. 27:1153–1158. Diekema, D. J., et al. 1999. Survey of bloodstream infections due to gramnegative bacilli: frequency of occurrence and antimicrobial susceptibility of isolates collected in the United States, Canada, and Latin America for the SENTRY Antimicrobial Surveillance Program, 1997. Clin. Infect. Dis. 29: 595–607. Dodson, W. H. 1968. Serratia marcescens septicemia. Arch. Intern. Med. 121:145–150. Doi, Y., B.-S. Lee, R. Iriye, S. Tabata, and K. Tateishi. 1998. Dominantly growing bacteria in malodorless domestic sewage treatment tanks and their biochemical characteristics. J. Antibact. Antifungal Agents (Japan) 26: 53–63. Doi, Y., et al. 2004. Plasmid-mediated 16S rRNA methylase in Serratia marcescens conferring high-level resistance to aminoglycosides. Antimicrob. Agents Chemother. 48:491–496. Domingo, D., et al. 1994. Nosocomial septicemia caused by Serratia plymuthica. J. Clin. Microbiol. 32:575–577. Donowitz, L. G., F. J. Marsik, J. W. Hoyt, and R. P. Wenzel. 1979. Serratia marcescens bacteremia from contaminated pressure inducers. JAMA 242: 1749–1751. Dorsey, G., et al. 2000. A heterogeneous outbreak of Enterobacter cloacae and Serratia marcescens infections in a surgical intensive care unit. Infect. Control Hosp. Epidemiol. 21:465–469. Dowzicky, M. J., and C. H. Park. 2008. Update on antimicrobial susceptibility rates among gram-negative and gram-positive organisms in the United States: results from the Tigecycline Evaluation and Surveillance Trial (TEST) 2005 to 2007. Clin. Ther. 30:2040–2050. Dubouix, A., et al. 2005. Epidemiological investigation of a Serratia liquefaciens outbreak in a neurosurgery department. J. Hosp. Infect. 60:8–13. Duggan, T. G., R. A. Leng, B. M. Hancock, and R. T. Cursons. 1984. Serratia marcescens in a newborn unit—microbiological features. Pathology 16:189– 191. Duncan, K. L., J. Ransley, and M. Elterman. 1994. Transfusion-transmitted Serratia liquefaciens from an autologous blood unit. Transfusion 34:738– 739. Dye, D. W. 1966. A comparative study of some atypical “xanthomonads.” N. Z. J. Sci. 9:843–854. Eberl, L., et al. 1996. Involvement of N-acyl-L-homoserine lactone autoinducers in controlling the multicellular behaviour of Serratia liquefaciens. Mol. Microbiol. 20:127–136. Echols, R. M., D. L. Palmer, R. M. King, and G. W. Long. 1984. Multidrugresistant Serratia marcescens bacteriuria related to urologic instrumentation. South. Med. J. 77:173–177. Ehrenkranz, N. J., E. A. Bolyard, M. Wiener, and T. J. Cleary. 1980. Antibiotic-sensitive Serratia marcescens infections complicating cardiopulmonary operations: contaminated disinfectant as a reservoir. Lancet 316: 1289–1292. Empel, J., et al. 2008. Molecular survey of ␤-lactamases conferring resistance to newer ␤-lactams in Enterobacteriaceae isolates from Polish hospitals. Antimicrob. Agents Chemother. 52:2449–2454. Engelhart, S., et al. 2003. Severe Serratia liquefaciens sepsis following vitamin C infusion treatment by a naturopathic practitioner. J. Clin. Microbiol. 41:3986–3988. Esel, D., et al. 2002. Polymicrobial ventriculitis and evaluation of an out-

786

125. 126.

127.

128.

129. 130. 131.

132. 133. 134.

135.

136.

137.

138.

139. 140. 141.

142.

143.

144. 145. 146.

147.

148.

149.

150.

151.

152.

MAHLEN break in a surgical intensive care unit due to inadequate sterilization. J. Hosp. Infect. 50:170–174. Ewing, W. H. 1963. An outline of nomenclature for the family Enterobacteriaceae. Int. Bull. Bacteriol. Nomencl. Taxon. 13:95–110. Ewing, W. H., B. R. Davis, M. A. Fife, and E. F. Lessel. 1973. Biochemical characterization of Serratia liquefaciens (Grimes and Hennerty) Bascomb et al. (formerly Enterobacter liquefaciens) and Serratia rubidaea (Stapp) comb. nov. and designation of type and neotype strains. Int. J. Syst. Bacteriol. 23:217–225. Ewing, W. H., B. R. Davis, and R. W. Reavis. 1959. Studies on the Serratia group. In CDC laboratory manual. Communicable Disease Center, CDC, Atlanta, GA. Farmer, J. J., III, K. D. Boatwright, and J. M. Janda. 2007. Enterobacteriaceae: introduction and identification, p. 649–669. In P. R. Murray, E. J. Baron, J. H. Jorgensen, M. L. Landry, and M. A. Pfaller (ed.), Manual of clinical microbiology, 9th ed. ASM Press, Washington, DC. Farmer, J. J., III, B. R. Davis, P. A. D. Grimont, and F. Grimont. 1977. Source of American Serratia. Lancet 310:459–460. Farmer, J. J., III, et al. 1976. Detection of Serratia outbreaks in hospital. Lancet 308:455–459. Farmer, J. J., III, et al. 1985. Biochemical identification of new species and biogroups of Enterobacteriaceae isolated from clinical specimens. J. Clin. Microbiol. 21:46–76. Fitzgerald, P., J. H. Drew, and I. Kruszelnicki. 1984. Serratia: a problem in a neonatal nursery. Aust. Paediatr. J. 20:205–207. Fleisch, F., et al. 2002. Three consecutive outbreaks of Serratia marcescens in a neonatal intensive care unit. Clin. Infect. Dis. 34:767–773. Fontana, R., et al. 2001. Antimicrobial susceptibility of respiratory isolates of Enterobacteriaceae and Staphylococcus aureus in Italy: incidence and trends over the period 1997–1999. Eur. J. Clin. Microbiol. Infect. Dis. 20:854–863. Franczek, S. P., R. P. Williams, and S. I. Hull. 1986. A survey of potential virulence factors in clinical and environmental isolates of Serratia marcescens. J. Med. Microbiol. 22:151–156. Frean, J. A., L. Arntzen, I. Rosekilly, and M. Isaa ¨cson. 1994. Investigation of contaminated parenteral nutrition fluids associated with an outbreak of Serratia odorifera septicaemia. J. Hosp. Infect. 27:263–273. Friedman, N. D., D. Kotsanas, J. Brett, B. Billah, and T. M. Korman. 2008. Investigation of an outbreak of Serratia marcescens in a neonatal unit via a case-control study and molecular typing. Am. J. Infect. Control 36:22–28. Fritsche, T. R., P. A. Strabala, H. S. Sader, M. J. Dowzicky, and R. N. Jones. 2005. Activity of tigecycline tested against a global collection of Enterobacteriaceae, including tetracycline-resistant isolates. Diagn. Microbiol. Infect. Dis. 52:209–213. Gale, D., and J. D. Lord. 1957. Overgrowth of Serratia marcescens in respiratory tract, simulating hemoptysis: report of a case. JAMA 164:1328–1331. Gale, D., and A. C. Sonnenwirth. 1962. Frequent human isolations of Serratia marcescens. Arch. Intern. Med. 109:414–421. Gallois, A., and P. A. D. Grimont. 1985. Pyrazines responsible for the potatolike odor produced by some Serratia and Cedecia strains. Appl. Environ. Microbiol. 50:1048–1051. Gargallo-Viola, D. 1989. Enzyme polymorphism, prodigiosin production, and plasmid fingerprints in clinical and naturally occurring isolates of Serratia marcescens. J. Clin. Microbiol. 27:860–868. Garza-Gonza ´lez, E., S. I. Mendoza Ibarra, J. M. Llaca-Díaz, and G. M. Gonzalez. 2011. Molecular characterization and antimicrobial susceptibility of extended-spectrum ␤-lactamase-producing Enterobacteriaceae isolates at a tertiary-care centre in Monterrey, Mexico. J. Med. Microbiol. 60:84–90. Gaughran, E. R. L. 1969. From superstition to science: the history of a bacterium. Trans. N. Y. Acad. Sci. 31:3–24. Gavini, F., et al. 1979. Serratia fonticola, a new species from water. Int. J. Syst. Bacteriol. 29:92–101. Geiger, A., M.-L. Fardeau, E. Falsen, B. Ollivier, and G. Cuny. 2010. Serratia glossinae sp. nov., isolated from the midgut of the tsetse fly Glossina palpalis gambiensis. Int. J. Syst. Evol. Microbiol. 60:1261–1265. Geiseler, P. J., B. Harris, and B. R. Andersen. 1982. Nosocomial outbreak of nitrate-negative Serratia marcescens infections. J. Clin. Microbiol. 15: 728–730. Giles, M., et al. 2006. What is the best screening method to detect Serratia marcescens colonization during an outbreak in a neonatal intensive care nursery? J. Hosp. Infect. 62:349–352. Gill, V. J., J. J. Farmer III, P. A. D. Grimont, M. A. Asbury, and C. L. McIntosh. 1981. Serratia ficaria isolated from a human clinical specimen. J. Clin. Microbiol. 14:234–236. Gillespie, E. E., J. Bradford, J. Brett, and D. Kotsanas. 2007. Serratia marcescens bacteremia—an indicator for outbreak management and heightened surveillance. J. Perinat. Med. 35:227–231. Glare, T. R., G. E. Corbett, and T. J. Sadler. 1993. Association of a large plasmid with amber disease of the New Zealand grass grub, Costelytra zealandica, caused by Serratia entomophila and Serratia proteamaculans. J. Invertebr. Pathol. 62:165–170. Glustein, J. Z., B. Rudensky, and A. Abrahamov. 1994. Catheter-associated

CLIN. MICROBIOL. REV.

153. 154. 155.

156.

157.

158.

159.

160.

161. 162. 163.

164.

165.

166.

167.

168.

169.

170.

171.

172. 173.

174. 175.

176. 177. 178.

179. 180.

sepsis caused by Serratia odorifera biovar 1 in an adolescent patient. Eur. J. Clin. Microbiol. Infect. Dis. 13:183–184. Goluszko, P., and M. R. Nowacki. 1989. Extracellular haemolytic activity of Serratia marcescens. FEMS Microbiol. Lett. 61:207–211. Gorret, J., et al. 2009. Childhood delayed septic arthritis of the knee caused by Serratia fonticola. Knee 16:512–514. Goullet, P., and B. Picard. 1997. An epidemiological study of Serratia marcescens isolates from nosocomial infections by enzyme electrophoresis. J. Med. Microbiol. 46:1019–1028. Gransden, W. R., M. Webster, G. L. French, and I. Phillips. 1986. An outbreak of Serratia marcescens transmitted by contaminated breast pumps in a special care baby unit. J. Hosp. Infect. 7:149–154. Grauel, E. L., E. Halle, R. Bollmann, P. Buchholz, and S. Buttenberg. 1989. Neonatal septicaemia—incidence, etiology, and outcome. A 6-year analysis. Acta Paediatr. Scand. Suppl. 360:113–119. Grimes, M. 1961. Classification of the Klebsiella-Aerobacter group with special reference to the cold-tolerant mesophilic Aerobacter types. Int. Bull. Bacteriol. Nomencl. Taxon. 11:111–129. Grimont, F., and P. A. D. Grimont. 2005. Genus XXXIV. Serratia Bizio 1823, 288AL, p. 799–811. In D. J. Brenner, N. R. Krieg, and J. T. Staley (ed.), Bergey’s manual of systematic bacteriology, 2nd ed., vol. 2, part B. Springer Science and Business Media, New York, NY. Grimont, F., and P. A. D. Grimont. 2006. The genus Serratia, p. 219–244. In M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer, and E. Stackebrandt (ed.), The prokaryotes, 3rd ed., vol. 6. Springer Science and Business Media, New York, NY. Grimont, P. A. D., and F. Grimont. 1978. The genus Serratia. Annu. Rev. Microbiol. 32:221–248. Grimont, P. A. D., F. Grimont, and H. L. C. Dulong de Rosney. 1977. Taxonomy of the genus Serratia. J. Gen. Microbiol. 98:39–66. Grimont, P. A. D., F. Grimont, and K. Irino. 1983. Biochemical characterization of Serratia liquefaciens sensu stricto, Serratia proteamaculans, and Serratia grimesii sp. nov. Curr. Microbiol. 7:69–74. Grimont, P. A. D., F. Grimont, and O. Lysenko. 1979. Species and biotype identification of Serratia strains associated with insects. Curr. Microbiol. 2:139–142. Grimont, P. A. D., et al. 1978. Deoxyribonucleic acid relatedness between Serratia plymuthica and other Serratia species, with a description of Serratia odorifera sp. nov. (type strain: ICPB 3995). Int. J. Syst. Bacteriol. 28:453– 463. Grimont, P. A. D., F. Grimont, and M. P. Starr. 1978. Serratia proteamaculans (Paine and Stansfield) comb. nov., a senior subjective synonym of Serratia liquefaciens (Grimes and Hennerty) Bascomb et al. Int. J. Syst. Bacteriol. 28:503–510. Grimont, P. A. D., F. Grimont, and M. P. Starr. 1979. Serratia ficaria sp. nov., a bacterial species associated with Smyrna figs and the fig wasp Blastophaga psenes. Curr. Microbiol. 2:277–282. Grimont, P. A. D., F. Grimont, and M. P. Starr. 1981. Comment on the request to the judicial commission to conserve the specific epithet liquefaciens over the specific epithet proteamaculans in the name of the organism currently known as Serratia liquefaciens. Int. J. Syst. Bacteriol. 31:211–212. Grimont, P. A. D., T. A. Jackson, E. Ageron, and M. J. Noonan. 1988. Serratia entomophila sp. nov. associated with amber disease in the New Zealand grass grub Costelytra zealandica. Int. J. Syst. Bacteriol. 38:1–6. Grkovic, S., T. R. Glare, T. A. Jackson, and G. E. Corbett. 1995. Genes essential for amber disease in grass grubs are located on the large plasmid found in Serratia entomophila and Serratia proteamaculans. Appl. Environ. Microbiol. 61:2218–2223. Grohskopf, L. A., et al. 2001. Serratia liquefaciens bloodstream infections from contamination of epoetin alfa at a hemodialysis center. N. Engl. J. Med. 344:1491–1497. Gurevitch, J., and D. Weber. 1950. A strain of Serratia isolated from urine. Am. J. Clin. Pathol. 20:48–49. Hanson, N. D., and C. C. Sanders. 1999. Regulation of inducible AmpC beta-lactamase expression among Enterobacteriaceae. Curr. Pharm. Des. 5:881–894. Harden, W. B., J. F. Fisher, D. H. Loebl, and J. P. Bailey. 1980. Septic arthritis due to Serratia liquefaciens. Arthritis Rheum. 23:946–947. Harnett, S. J., K. D. Allen, and R. R. Macmillan. 2001. Critical care unit outbreak of Serratia liquefaciens from contaminated pressure monitoring equipment. J. Hosp. Infect. 47:301–307. Harrison, F. C. 1924. The “miraculous” micro-organism. Trans. R. Soc. Can. 18:1–17. Hawe, A. J., and M. H. Hughes. 1954. Bacterial endocarditis due to Chromobacterium prodigiosum. Br. Med. J. 1:968–970. Hejazi, A., H. M. Aucken, and F. R. Falkiner. 2000. Epidemiology and susceptibility of Serratia marcescens in a large general hospital over an 8-year period. J. Hosp. Infect. 45:42–46. Hejazi, A., and F. R. Falkiner. 1997. Serratia marcescens. J. Med. Microbiol. 46:903–912. Henriques, I., A. Moura, A. Alves, M. J. Saavedra, and A. Correia. 2004. Molecular characterization of a carbapenem-hydrolyzing class A beta-lac-

VOL. 24, 2011

181.

182.

183. 184. 185.

186.

187.

188.

189. 190.

191. 192. 193. 194.

195.

196. 197. 198.

199.

200.

201. 202.

203. 204. 205.

206. 207. 208.

209.

210.

tamase, SFC-1, from Serratia fonticola UTAD54. Antimicrob. Agents Chemother. 48:2321–2324. Henry, B., C. Plante-Jenkins, and K. Ostrowska. 2001. An outbreak of Serratia marcescens associated with the anesthetic agent propofol. Am. J. Infect. Control 29:312–315. Herbert, S., et al. 2007. Large outbreak of infection and colonization with gram-negative pathogens carrying the metallo-␤-lactamase gene blaIMP-4 at a 320-bed tertiary hospital in Australia. Infect. Control Hosp. Epidemiol. 28:98–101. Herra, C. M., et al. 1998. An outbreak of an unusual strain of Serratia marcescens in two Dublin hospitals. J. Hosp. Infect. 39:135–141. Hertle, R. 2005. The family of Serratia type pore forming toxins. Curr. Protein Pept. Sci. 6:313–325. Holmes, B., and J. J. Farmer III. 2009. International Committee on Systematics of Prokaryotes Subcommittee on the Taxonomy of Enterobacteriaceae. Minutes of the meetings, 7 August 2008, Instanbul, Turkey. Int. J. Syst. Evol. Microbiol. 59:2643–2645. Horcajada, J. P., et al. 2006. Acquisition of multidrug-resistant Serratia marcescens by critically ill patients who consumed tap water during receipt of oral medication. Infect. Control Hosp. Epidemiol. 27:774–777. Horng, Y. T., et al. 2002. The LuxR family protein SpnR functions as a negative regulator of N-acylhomoserine lactone-dependent quorum sensing in Serratia marcescens. Mol. Microbiol. 45:1655–1671. Hornsey, M., et al. 2010. Tigecycline resistance in Serratia marcescens associated with up-regulation of the SdeXY-HasF efflux system also active against ciprofloxacin and cefpirome. J. Antimicrob. Chemother. 65:479– 482. Horowitz, H. W., et al. 1987. Serratia plymuthica sepsis associated with infection of central venous catheter. J. Clin. Microbiol. 25:1562–1563. Hoyen, C., et al. 1999. Use of real time pulsed field gel electrophoresis to guide interventions during a nursery outbreak of Serratia marcescens infection. Pediatr. Infect. Dis. J. 18:357–360. Hugh-Jones, M. 1992. Wickham Steed and German biological warfare research. Intell. Natl. Secur. 7:379–402. Huovinen, P. 2001. Resistance to trimethoprim-sulfamethoxazole. Clin. Infect. Dis. 32:1608–1614. Hurrell, E., et al. 2009. Neonatal enteral feeding tubes as loci for colonisation by members of the Enterobacteriaceae. BMC Infect. Dis. 9:146. Ishikawa, K., et al. 2011. The nationwide study of bacterial pathogens associated with urinary tract infections conducted by the Japanese Society of Chemotherapy. J. Infect. Chemother. 17:126–138. Itami, C., H. Matsunaga, T. Sawada, H. Sakurai, and Y. Kimura. 1993. Superoxide dismutase mimetic activities of metal complexes of L-2(2-pyridyl)-1-pyrroline-5-carboxylic acid (pyrimine). Biochem. Biophys. Res. Commun. 197:536–541. Ivanova, D., et al. 2008. Extended-spectrum ␤-lactamase-producing Serratia marcescens outbreak in a Bulgarian hospital. J. Hosp. Infect. 70:60–65. Jacoby, G. A. 2009. AmpC ␤-lactamases. Clin. Microbiol. Rev. 22:161–182. Jang, T. N., et al. 2001. Use of pulsed-field gel electrophoresis to investigate an outbreak of Serratia marcescens infection in a neonatal intensive care unit. J. Hosp. Infect. 48:13–19. Jean, S. S., et al. 2009. Nationwide surveillance of antimicrobial resistance among Enterobacteriaceae in intensive care units in Taiwan. Eur. J. Clin. Microbiol. Infect. Dis. 28:215–220. Jeppsson, B., S. Lindahl, S. Ingemansson, S. Kornhall, and S. Sjovall. 1984. Bacterial contamination of blood transfusion: an unusual cause of sepsis. Acta Chir. Scand. 150:489–491. Jepson, A. P., et al. 2006. Finger rings should be removed prior to scrubbing. J. Hosp. Infect. 64:197–198. John, J. F., Jr., and W. F. McNeill. 1981. Characteristics of Serratia marcescens containing a plasmid coding for gentamicin resistance in nosocomial infections. J. Infect. Dis. 143:810–817. Johnson, E., and P. D. Ellner. 1974. Distribution of Serratia species in clinical specimens. Appl. Microbiol. 28:513–514. Jones, B. L., et al. 2000. An outbreak of Serratia marcescens in two neonatal intensive care units. J. Hosp. Infect. 46:314–319. Jones, R. N. 2010. Microbial etiologies of hospital-acquired bacterial pneumonia and ventilator-associated bacterial pneumonia. Clin. Infect. Dis. 51(Suppl. 1):S81–S87. Jones, S. R., M. Amon, C. Falvey, and K. Patrick. 1978. Serratia marcescens colonizing the gut. Lancet 311:1105. Joondeph, H. C., and A. F. Nothnagel. 1983. Serratia rubidae endophthalmitis following penetrating ocular injury. Ann. Ophthalmol. 15:1138–1140. Kalbe, C., P. Marten, and G. Berg. 1996. Strains of the genus Serratia as beneficial rhizobacteria of oilseed rape with antifungal properties. Microbiol. Res. 151:433–439. Kanagasabhapathy, M., G. Yamazaki, A. Ishida, H. Sasaki, and S. Nagata. 2009. Presence of quorum-sensing inhibitor-like compounds from bacteria isolated from the brown alga Colpomenia sinuosa. Lett. Appl. Microbiol. 49:573–579. Kang, H. Y., et al. 2008. Distribution of conjugative-plasmid-mediated 16S rRNA methylase genes among amikacin-resistant Enterobacteriaceae iso-

SERRATIA INFECTIONS

211.

212.

213.

214.

215.

216.

217.

218.

219.

220. 221.

222.

223.

224.

225.

226.

227.

228.

229.

230.

231. 232.

233.

234.

787

lates collected in 1995 to 1998 and 2001 to 2006 at a university hospital in South Korea and identification of conjugative plasmids mediating dissemination of 16S rRNA methylase. J. Clin. Microbiol. 46:700–706. Kappstein, I., C. M. Schneider, H. Grundmann, R. Scholz, and P. Janknecht. 1999. Long-lasting contamination of a vitrectomy apparatus with Serratia marcescens. Infect. Control Hosp. Epidemiol. 20:192–195. Kass, E. H., and L. J. Schneiderman. 1957. Entry of bacteria into the urinary tracts of patients with inlying catheters. N. Engl. J. Med. 256:556– 557. Kim, B.-N., S.-I. Choi, and N.-H. Ryoo. 2006. Three-year follow-up of an outbreak of Serratia marcescens bacteriuria in a neurosurgical intensive care unit. J. Korean Med. Sci. 21:973–978. Kim, J., and Y.-M. Lim. 2005. Prevalence of derepressed AmpC mutants and extended-spectrum ␤-lactamase producers among clinical isolates of Citrobacter freundii, Enterobacter spp., and Serratia marcescens in Korea: dissemination of CTX-M-3, TEM-52, and SHV-12. J. Clin. Microbiol. 43: 2452–2455. Kim, J.-H., W.-H. Choi, S.-W. Yun, S.-A. Chae, and B.-H. Yoo. 2010. An outbreak of Serratia marcescens sepsis in a pediatric ward. Clin. Pediatr. 49:1000–1002. Kim, J. H., E. H. Cho, K. S. Kim, H. Y. Kim, and Y. M. Kim. 1998. Cloning and nucleotide sequence of the DNA gyrase gyrA gene from Serratia marcescens and characterization of mutations in gyrA of quinolone-resistant clinical isolates. Antimicrob. Agents Chemother. 42:190–193. Kim, S.-Y., Y.-J. Park, J. K. Yu, Y. S. Kim, and K. Han. 2009. Prevalence and characteristics of aac(6⬘)-Ib-cr in AmpC-producing Enterobacter cloacae, Citrobacter freundii, and Serratia marcescens: a multicenter study from Korea. Diagn. Microbiol. Infect. Dis. 63:314–318. Kiratisin, P., and A. Henprasert. 2011. Resistance phenotype-genotype correlation and molecular epidemiology of Citrobacter, Enterobacter, Proteus, Providencia, Salmonella and Serratia that carry extended-spectrum ␤-lactamases with or without plasmid-mediated AmpC ␤-lactamase genes in Thailand. Trans. R. Soc. Trop. Med. Hyg. 105:46–51. Kleef, R., and E. D. Hager. 2006. Fever, pyrogens and cancer, p. 276–337. In G. F. Baronzio and E. D. Hager (ed.), Hyperthermia in cancer treatment: a primer. Springer Science, New York, NY. Klein, E. 1894. On an infection of food-stuffs by Bacillus prodigiosus. J. Pathol. Bacteriol. 2:217–218. Knowles, S., et al. 2000. An outbreak of multiply resistant Serratia marcescens: the importance of persistent carriage. Bone Marrow Transplant. 25:873–877. Krieger, J. N., E. Levy-Zombek, A. Scheidt, and L. M. Drusin. 1980. A nosocomial epidemic of antibiotic-resistant Serratia marcescens urinary tract infections. J. Urol. 124:498–502. Krishnan, P. U., B. Pereira, and R. Macaden. 1991. Epidemiological study of an outbreak of Serratia marcescens in a haemodialysis unit. J. Hosp. Infect. 18:57–61. Kumar, A., and E. A. Worobec. 2005. Cloning, sequencing, and characterization of the SdeAB multidrug efflux pump of Serratia marcescens. Antimicrob. Agents Chemother. 49:1495–1501. Kunimoto, D., R. Rennie, D. M. Citron, and E. J. C. Goldstein. 2004. Bacteriology of a bear bite wound to a human: case report. J. Clin. Microbiol. 42:3374–3376. Kurz, C. L., et al. 2003. Virulence factors of the human opportunistic pathogen Serratia marcescens identified by in vivo screening. EMBO J. 22:1451–1460. Lai, H. C., et al. 2005. The RssAB two-component signal transduction system in Serratia marcescens regulates swarming motility and cell envelope architecture in response to exogenous saturated fatty acids. J. Bacteriol. 187:3407–3414. Lai, K. K., S. P. Baker, and S. A. Fontecchio. 2004. Rapid eradication of a cluster of Serratia marcescens in a neonatal intensive care unit: use of epidemiologic chromosome profiling by pulsed-field gel electrophoresis. Infect. Control Hosp. Epidemiol. 25:730–734. Lakso, J. U., and M. P. Starr. 1970. Comparative injuriousness to plants of Erwinia spp. and other enterobacteria from plants and animals. J. Appl. Bacteriol. 33:692–707. Lamelas, A., et al. 2008. Evolution of the secondary symbiont “Candidatus Serratia symbiotica” in aphid species of the subfamily Lachninae. Appl. Environ. Microbiol. 74:4236–4240. Lancaster, L. J. 1962. Role of Serratia species in urinary tract infections. Arch. Intern. Med. 109:536–539. Lauderdale, T. L., et al. 2004. The status of antimicrobial resistance in Taiwan among gram-negative pathogens: the Taiwan surveillance of antimicrobial resistance (TSAR) program, 2000. Diagn. Microbiol. Infect. Dis. 48:211–219. Laupland, K. B., et al. 2008. Population-based laboratory surveillance for Serratia species isolates in a large Canadian health region. Eur. J. Clin. Microbiol. Infect. Dis. 27:89–95. Lazachek, G. W., G. L. Boyle, A. L. Schwartz, and I. H. Leopold. 1971. Serratia marcescens, an ocular pathogen. New considerations. Arch. Ophthalmol. 86:599–603.

788

MAHLEN

235. Lee, J., J. Carey, and D. C. Perlman. 2006. Pneumonia and bacteremia due to Serratia odorifera. J. Infect. 53:212–214. 236. Lelliott, R. A. 1974. Genus XII. Erwinia Winslow, Broadhurst, Buchanon, Krumwiede, Rogers and Smith 1920, 209, p. 332–339. In R. E. Buchanon and N. E. Gibbons (ed.), Bergey’s manual of determinative bacteriology, 8th ed. The Williams and Wilkins Co., Baltimore, MD. 237. Lewis, A. M., J. R. Stephenson, J. Garner, F. Afshar, and S. Tabaqchali. 1989. A hospital outbreak of Serratia marcescens in neurosurgical patients. Epidemiol. Infect. 102:69–74. 238. Lewis, D. A., et al. 1983. Infection with netilmicin resistant Serratia marcescens in a special care baby unit. Br. Med. J. 287:1701–1705. 239. Ligozzi, M., R. Fontana, M. Aldegheri, G. Scalet, and G. Lo Cascio. 2010. Comparative evaluation of an automated repetitive-sequence based PCR instrument versus pulsed-field gel electrophoresis in the setting of a Serratia marcescens nosocomial infection outbreak. J. Clin. Microbiol. 48:1690– 1695. 240. Lin, C. S., et al. 2010. RssAB-FlhDC-ShlBA as a major pathogenesis pathway in Serratia marcescens. Infect. Immun. 78:4870–4881. 241. Liu, D., et al. 2010. Outbreak of Serratia marcescens infection due to contamination of multiple-dose vial of heparin-saline solution used to flush deep venous catheters or peripheral trocars. J. Hosp. Infect. 77:175–176. 242. Liu, P. Y., et al. 1994. Use of PCR to study epidemiology of Serratia marcescens isolates in nosocomial infection. J. Clin. Microbiol. 32:1935– 1938. 243. Liu, X., et al. 2011. Characterisation of two quorum sensing systems in the endophytic Serratia plymuthica strain G3: differential control of motility and biofilm formation according to life-style. BMC Microbiol. 11:26. 244. Livermore, D. M., T. G. Winstanley, and K. P. Shannon. 2001. Interpretative reading: recognizing the unusual and inferring resistance mechanisms from resistance phenotypes. J. Antimicrob. Chemother. 48(Suppl. 1):87– 102. 245. Lockhart, S. R., et al. 2007. Antimicrobial resistance among gram-negative bacilli causing infections in intensive care unit patients in the United States between 1993 and 2004. J. Clin. Microbiol. 45:3352–3359. 246. Luzzaro, F., et al. 1998. Repeated epidemics caused by extended-spectrum beta-lactamase-producing Serratia marcescens strains. Eur. J. Clin. Microbiol. Infect. Dis. 17:629–636. 247. Madduri, S. D., D. A. Mauriello, L. G. Smith, and J. J. Seebode. 1976. Serratia marcescens and the urologist. J. Urol. 116:613–615. 248. Mahlen, S. D., S. S. Morrow, B. Abdalhamid, and N. D. Hanson. 2003. Analyses of ampC gene expression in Serratia marcescens reveal new regulatory properties. J. Antimicrob. Chemother. 51:791–802. 249. Manning, M. L., L. K. Archibald, L. M. Bell, S. J. Banerjee, and W. R. Jarvis. 2001. Serratia marcescens transmission in a pediatric intensive care unit: a multifactorial occurrence. Am. J. Infect. Control 29:115–119. 250. Maragakis, L. L., et al. 2008. Outbreak of multi-drug resistant Serratia marcescens infection in a neonatal intensive care unit. Infect. Control Hosp. Epidemiol. 29:418–423. 251. Marrie, T. J., and J. W. Costerton. 1981. Prolonged survival of Serratia marcescens in chlorhexidine. Appl. Environ. Microbiol. 42:1093–1102. 252. Martincic, I., C. Mastronardi, A. Chung, and S. Ramirez-Arcos. 2008. Unexplained agglutination of stored red blood cells in Alsever’s solution caused by the gram-negative bacterium Serratia liquefaciens. Immunohematology 24:39–44. 253. Martinez-Martinez, L., A. Pascual, and G. A. Jacoby. 1998. Quinolone resistance from a transferable plasmid. Lancet 351:797–799. 254. Masecar, B. L., and N. J. Robillard. 1991. Spontaneous quinolone resistance in Serratia marcescens due to a mutation in gyrA. Antimicrob. Agents Chemother. 35:898–902. 255. Maseda, H., Y. Hashida, R. Konaka, A. Shirai, and H. Kourai. 2009. Mutational upregulation of a resistance-nodulation-cell division-type multidrug efflux pump, SdeAB, upon exposure to a biocide, cetylpyridinium chloride, and antibiotic resistance in Serratia marcescens. Antimicrob. Agents Chemother. 53:5230–5235. 256. Matsumoto, K. 2004. Role of bacterial proteases in pseudomonal and serratial keratitis. Biol. Chem. 385:1007–1016. 257. Matsuo, T., et al. 2008. SmdAB, a heterodimeric ABC-type multidrug efflux pump, in Serratia marcescens. J. Bacteriol. 190:648–654. 258. McAllister, T. A., et al. 1989. Serratia marcescens outbreak in a paediatric oncology unit traced to contaminated chlorhexidine. Scott. Med. J. 34:525– 528. 259. McCormack, R. C., and C. M. Kunin. 1966. Control of a single source nursery epidemic due to Serratia marcescens. Pediatrics 37:750–755. 260. McEntegart, M. G., and J. S. Porterfield. 1949. Bacteraemia following dental extractions. Lancet 254:596–598. 261. Mellow, M. H., and R. J. Lewis. 1976. Endoscopy-related bacteremia. Incidence of positive blood cultures after endoscopy of upper gastrointestinal tract. Arch. Intern. Med. 136:667–669. 262. Mendonc¸a-Caridad, J. J., P. Juiz Lopez, L. Francos, and M. Rodriguez. 2008. Swordfish bill injury involving the pterygomaxillary fossae: surgical management and case report. J. Oral Maxillofac. Surg. 66:1739–1743. 263. Menezes, E. A., F. C. Cezafar, S. Andrade Mdo, M. V. Rocha, and F. A.

CLIN. MICROBIOL. REV.

264.

265. 266. 267.

268. 269.

270.

271.

272.

273.

274. 275.

276.

277. 278. 279.

280. 281.

282.

283.

284.

285.

286. 287.

288.

289. 290.

291.

Cunha. 2004. Frequency of Serratia sp. in urine infections of intern patients in the Santa Casa de Miserico ´rdia in Fortaleza. Rev. Soc. Bras. Med. Trop. 37:70–71. (In Portuguese.) Merlino, C. P. 1924. Bartolomeo Bizio’s letter to the most eminent priest, Angelo Bellani, concerning the phenomenon of the red-colored polenta. J. Bacteriol. 9:527–543. Mermel, L. A., and C. A. Spiegel. 1992. Nosocomial sepsis due to Serratia odorifera biovar 1. Clin. Infect. Dis. 14:208–210. Midelfart, J., A. Midelfart, and L. Bevanger. 1996. Microbial contamination of contact lens cases among medical students. CLAO J. 22:21–24. Milisavljevic, V., et al. 2004. Molecular epidemiology of Serratia marcescens outbreaks in two neonatal intensive care units. Infect. Control Hosp. Epidemiol. 25:719–721. Mills, J., and D. Drew. 1976. Serratia marcescens endocarditis: a regional illness associated with intravenous drug abuse. Ann. Intern. Med. 84:29–35. Miranda, G., et al. 1996. Use of pulsed-field gel electrophoresis typing to study an outbreak of infection due to Serratia marcescens in a neonatal intensive care unit. J. Clin. Microbiol. 34:3138–3141. Miranda-Novales, G., et al. 2003. An outbreak due to Serratia marcescens in a neonatal intensive care unit typed by 2-day pulsed-field gel electrophoresis protocol. Arch. Med. Res. 34:237–241. Miriagou, V., et al. 2008. Emergence of Serratia liquefaciens and Klebsiella oxytoca with metallo-␤-lactamase-encoding IncW plasmids: further spread of the blaVIM-1-carrying integron In-e541. Int. J. Antimicrob. Agents 32: 540–541. Mlynarczyk, A., et al. 2007. Serratia marcescens isolated in 2005 from clinical specimens from patients with diminished immunity. Transplant. Proc. 39:2879–2882. Mlynarczyk, A., et al. 2009. CTX-M and TEM as predominant types of extended spectrum ␤-lactamases among Serratia marcescens isolated from solid organ recipients. Transplant. Proc. 41:3253–3255. Moloney, A. C., A. H. Quoraishi, P. Parry, and V. Hall. 1987. A bacteriological examination of breast pumps. J. Hosp. Infect. 9:169–174. Montanaro, D., et al. 1984. Epidemiological and bacteriological investigation of Serratia marcescens epidemic in a nursery and in a neonatal intensive care unit. J. Hyg. (London) 93:67–78. Mossad, S. B. 2000. The world’s first case of Serratia liquefaciens intravascular catheter-related suppurative thrombophlebitis and native valve endocarditis. Clin. Microbiol. Infect. 6:559–560. Mostafa, E., et al. 2008. Septic pseudarthrosis caused by Serratia plymuthica. Joint Bone Spine 75:506–512. Muller, A. E., et al. 2010. Outbreak of severe sepsis due to contaminated propofol: lessons to learn. J. Hosp. Infect. 76:225–230. Mu ¨ller, H., et al. 2009. Quorum-sensing effects in the antagonistic rhizosphere bacterium Serratia plymuthica HRO-C48. FEMS Microbiol. Ecol. 67:468–478. Mutton, K. J., L. M. Brady, and J. L. Harkness. 1981. Serratia crossinfection in an intensive therapy unit. J. Hosp. Infect. 2:85–91. Nagy, E., Z. Pragai, Z. Ko ´czia ´n, E. Hajdu ´, and E. Fodor. 1998. Investigation of the presence of different broad-spectrum ␤-lactamases among clinical isolates of Enterobacteriaceae. Acta Microbiol. Immunol. Hung. 45:433–436. Nakashima, A. K., A. K. Highsmith, and W. J. Martone. 1987. Survival of Serratia marcescens in benzylalkonium chloride and in multiple-dose medication vials: relationship to epidemic septic arthritis. J. Clin. Microbiol. 25:1019–1021. Nakashima, A. K., M. A. McCarthy, W. J. Martone, and R. L. Anderson. 1987. Epidemic septic arthritis caused by Serratia marcescens and associated with a benzylalkonium chloride antiseptic. J. Clin. Microbiol. 25:1014–1018. Naumiuk, L., et al. 2004. Molecular epidemiology of Serratia marcescens in two hospitals in Gdan ´sk, Poland, over a 5-year period. J. Clin. Microbiol. 42:3108–3116. Newport, M. T., J. F. John, Y. M. Michel, and A. H. Levkoff. 1985. Endemic Serratia marcescens infection in a neonatal intensive care nursery associated with gastrointestinal colonization. Pediatr. Infect. Dis. 4:160–167. Nouh, F., and S. Bhandari. 2000. CAPD peritonitis due to Serratia plymuthica. Perit. Dial. Int. 20:349. Okuda, T., N. Endo, Y. Osada, and H. Zen-Yoji. 1984. Outbreak of nosocomial urinary tract infections caused by Serratia marcescens. J. Clin. Microbiol. 20:691–695. Osano, E., et al. 1994. Molecular characterization of an enterobacterial metallo beta-lactamase found in a clinical isolate of Serratia marcescens that shows imipenem resistance. Antimicrob. Agents Chemother. 38:71–78. Ostrowsky, B. E., et al. 2002. Serratia marcescens bacteremia traced to an infused narcotic. N. Engl. J. Med. 346:1529–1537. Overhage, J., M. Bains, M. D. Brazas, and R. E. Hancock. 2008. Swarming of Pseudomonas aeruginosa is a complex adaptation leading to increased production of virulence factors and antibiotic resistance. J. Bacteriol. 190: 2671–2679. Paine, S. G., and H. Stansfield. 1919. Studies in bacteriosis. III. A bacterial leaf-spot disease of Protea cynaroides, exhibiting a host reaction of possibly bacteriolytic nature. Ann. Appl. Biol. 6:27–39.

VOL. 24, 2011 292. Paine, T. F. 1946. Illness in man following inhalation of Serratia marcescens. J. Infect. Dis. 79:226–232. 293. Pałucha, A., B. Mikiewicz, W. Hryniewicz, and M. Gniadkowski. 1999. Concurrent outbreaks of extended-spectrum ␤-lactamase-producing organisms of the family Enterobacteriaceae in a Warsaw hospital. J. Antimicrob. Chemother. 44:489–499. 294. Papapanagiotou, J., and C. Aligizakis. 1959. Isolation of Chromobacterium prodigiosum from empyema. J. Clin. Pathol. 12:170–171. 295. Park, Y. J., et al. 2005. Occurrence of extended-spectrum beta-lactamases among chromosomal AmpC-producing Enterobacter cloacae, Citrobacter freundii, and Serratia marcescens in Korea and investigation of screening criteria. Diagn. Microbiol. Infect. Dis. 51:265–269. 296. Park, Y. J., J. K. Yu, S. Lee, E. J. Oh, and G. J. Woo. 2007. Prevalence and diversity of qnr alleles in AmpC-producing Enterobacter cloacae, Enterobacter aerogenes, Citrobacter freundii and Serratia marcescens: a multicentre study from Korea. J. Antimicrob. Chemother. 60:868–871. 297. Parvaz, P., et al. 2002. A rapid and easy PCR-RFLP method for genotyping Serratia marcescens strains isolated in different hospital outbreaks and patient environments in the Lyon area, France. J. Hosp. Infect. 51:96–105. 298. Pascual Pe´rez, R., et al. 2003. Necrotic cellulitis by Serratia plymuthica. Eur. J. Intern. Med. 14:501–503. 299. Passaro, D. J., et al. 1997. Postoperative Serratia marcescens wound infections traced to an out-of-hospital source. J. Infect. Dis. 175:992–995. 300. Paterson, D. L., and R. A. Bonomo. 2005. Extended-spectrum ␤-lactamases: a clinical update. Clin. Microbiol. Rev. 18:657–686. 301. Patterson, K. L., et al. 2002. The etiology of white pox, a lethal disease of the Caribbean elkhorn coral, Acropora palmata. Proc. Natl. Acad. Sci. U. S. A. 99:8725–8730. 302. Patterson, R. H., Jr., G. B. Banister, and V. Knight. 1952. Chromobacterial infection in man. AMA Arch. Intern. Med. 90:79–86. 303. Peleg, A. Y., C. Franklin, J. M. Bell, and D. W. Spelman. 2005. Dissemination of the metallo-␤-lactamase gene blaIMP-4 among gram-negative pathogens in a clinical setting in Australia. Clin. Infect. Dis. 41:1549–1556. 304. Peltroche-Llacsahuanga, H., R. Lu ¨tticken, and G. Haase. 1999. Temporally overlapping nosocomial outbreaks of Serratia marcescens infections: an unexpected result revealed by pulsed-field gel electrophoresis. Infect. Control Hosp. Epidemiol. 20:387–388. 305. Pfyffer, G. E. 1992. Serratia fonticola as an infectious agent. Eur. J. Clin. Microbiol. Infect. Dis. 11:299–300. 306. Picard, B., B. Bruneau, and P. Goullet. 1988. Demonstration of an outbreak of Serratia marcescens infections in a medical intensive care unit by esterase electrophoretic typing. J. Hosp. Infect. 11:194–195. 307. Pien, F. D., and J. J. Farmer III. 1983. Serratia ficaria isolated from a leg ulcer. South. Med. J. 76:1591–1592. 308. Pinna, A., D. Usai, L. A. Sechi, A. Carta, and S. Zanetti. 20 October 2009. Detection of virulence factors in Serratia strains isolated from contact lens-associated corneal ulcers. Acta Ophthalmol. doi:10.1111/j.17553768.2009.01689.x. 309. Prasad, G. A., P. G. Jones, J. Michaels, J. S. Garland, and C. R. Shivpuri. 2001. Outbreak of Serratia marcescens infection in a neonatal intensive care unit. Infect. Control Hosp. Epidemiol. 22:303–305. 310. Primavesi, R., D. A. Lewis, P. J. Fleming, and D. C. Speller. 1982. Serratia marcescens in a special baby unit. Lancet 317:1164. 311. Queenan, A. M., and K. Bush. 2007. Carbapenemases: the versatile ␤-lactamases. Clin. Microbiol. Rev. 20:440–458. 312. Queenan, A. M., et al. 2000. SME-type carbapenem-hydrolyzing class A beta-lactamases from geographically diverse Serratia marcescens strains. Antimicrob. Agents Chemother. 44:3035–3039. 313. Rabier, V., et al. 2008. Hand washing soap as a source of neonatal Serratia marcescens outbreak. Acta Paediatr. 97:1381–1385. 314. Rabinowitz, K., and R. Schiffrin. 1952. A ward contamination by Serratia marcescens. Acta Med. Orient. 11:181–184. 315. Ramírez-Arcos, S., et al. 2006. Fatal septic shock associated with transfusion-transmitted Serratia marcescens. Transfusion 46:679–680. 316. Ra ¨ndler, C., et al. 2010. A three-phase in-vitro system for studying Pseudomonas aeruginosa adhesion and biofilm formation upon hydrogel contact lenses. BMC Microbiol. 10:282. 317. Reina, J., N. Borell, and I. Lompart. 1992. Community-acquired bacteremia caused by Serratia plymuthica. Case report and review of the literature. Diagn. Microbiol. Infect. Dis. 15:449–452. 318. Rhomberg, P. R., and R. N. Jones. 2009. Summary trends for the Meropenem Yearly Susceptible Test information collection program: a 10-year experience in the United States (1999–2008). Diagn. Microbiol. Infect. Dis. 65:414–426. 319. Richards, N. M., and S. Levitsky. 1975. Outbreak of Serratia marcescens infections in a cardiothoracic surgicial intensive care unit. Ann. Thorac. Surg. 19:503–513. 320. Ringrose, R. E., et al. 1968. A hospital outbreak of Serratia marcescens associated with ultrasonic nebulizers. Ann. Intern. Med. 69:719–729. 321. Rizvi, M., et al. 2009. Extended spectrum AmpC and metallo-beta-lactamases in Serratia and Citrobacter spp. in a disc approximation assay. J. Infect. Dev. Ctries. 3:285–294.

SERRATIA INFECTIONS

789

322. Robertson, A. G., and L. J. Robertson. 1995. From asps to allegations: biological warfare in history. Mil. Med. 160:369–373. 323. Robicsek, A., et al. 2006. Fluoroquinolone-modifying enzyme: a new adaptation of a common aminoglycoside acetyltransferase. Nat. Med. 12:83–88. 324. Robinson, W., and P. B. Woolley. 1957. Pseudohaemoptysis due to Chromobacterium prodigiosum. Lancet 269:819. 325. Rodríguez-Martínez, J. M., M. E. Cano, C. Velasco, L. Martínez-Martínez, and A. Pascual. 2011. Plasmid-mediated quinolone resistance: an update. J. Infect. Chemother. 17:149–182. 326. Roth, V. R., et al. 2000. Transfusion-related sepsis due to Serratia liquefaciens in the United States. Transfusion 40:931–935. 327. Rudnick, J. R., et al. 1996. Gram-negative bacteremia in open-heart-surgery patients traced to probable tap-water contamination of pressure-monitoring equipment. Infect. Control Hosp. Epidemiol. 17:281–285. 328. Ruiz, N., T. Montero, J. Hernandez-Borrell, and M. Vin ˜ as. 2003. The role of Serratia marcescens porins in antibiotic resistance. Microb. Drug Resist. 9:257–264. 329. Rutala, W. A., V. A. Kennedy, H. B. Loflin, and F. A. Sarubbi, Jr. 1981. Serratia marcescens nosocomial infections of the urinary tract associated with urine measuring containers and urinometers. Am. J. Med. 70:659–663. 330. Sabri, A., et al. 5 November 2010. Isolation, pure culture and characterization of Serratia symbiotica, the R-type of secondary endosymbionts of the black bean aphid Aphis fabae. Int. J. Syst. Evol. Microbiol. doi:10.1099/ ijs.0.024133-0. 331. Sader, H. S., et al. 1994. Nosocomial transmission of Serratia odorifera biogroup 2: case report demonstration by macrorestriction analysis of chromosomal DNA using pulsed-field gel electrophoresis. Infect. Control Hosp. Epidemiol. 15:390–393. 332. Saito, H., L. Elting, G. P. Bodey, and P. Berkey. 1989. Serratia bacteremia: review of 118 cases. Rev. Infect. Dis. 11:912–920. 333. Samonis, G., et al. 10 January 2011. Serratia infections in a general hospital: characteristics and outcomes. Eur. J. Clin. Microbiol. Infect. Dis. doi: 10.1007/s10096-010-1135-4. 334. Sanders, C. C., and C. Watanakunakorn. 1986. Emergence of resistance to ␤-lactams, aminoglycosides, and quinolones during combination therapy for infection due to Serratia marcescens. J. Infect. Dis. 153:617–619. 335. Sanders, C. V., Jr., J. P. Luby, W. G. Johanson, J. A. Barnett, and J. P. Sanford. 1970. Serratia marcescens infections from inhalation therapy medications: nosocomial outbreak. Ann. Intern. Med. 73:15–21. 336. Sankaridurg, P. R., N. Vuppala, A. Sreedharan, J. Vadlamudi, and G. N. Rao. 1996. Gram negative bacteria and contact lens induced acute red eye. Indian J. Ophthalmol. 44:29–32. 337. Sartor, C., et al. 2000. Nosocomial Serratia marcescens infections associated with extrinsic contamination of a liquid nonmedicated soap. Infect. Control Hosp. Epidemiol. 21:196–199. 338. Sarvikivi, E., et al. 2004. Clustering of Serratia marcescens infections in a neonatal intensive care unit. Infect. Control Hosp. Epidemiol. 25:723–729. 339. Schaberg, D. R., et al. 1976. An outbreak of nosocomial infection due to multiply resistant Serratia marcescens: evidence of interhospital spread. J. Infect. Dis. 134:181–188. 340. Schaberg, D. R., R. A. Weinstein, and W. E. Stamm. 1976. Epidemics of nosocomial urinary tract infection caused by multiply resistant gram-negative bacilli: epidemiology and control. J. Infect. Dis. 133:363–366. 341. Sebert, M. E., M. L. Manning, K. L. McGowan, E. R. Alpern, and L. M. Bell. 2002. An outbreak of Serratia marcescens bacteremia after general anesthesia. Infect. Control Hosp. Epidemiol. 23:733–739. 342. Seeyave, D., N. Desai, S. Miller, S. P. Rao, and S. Piecuch. 2006. Fatal delayed transfusion reaction in a sickle cell anemia patient with Serratia marcescens sepsis. J. Natl. Med. Assoc. 98:1697–1699. 343. Sekhsokh, Y., et al. 2007. Serratia rubidaea bacteremia. Med. Mal. Infect. 37:287–289. (In French.) 344. Serruys-Schoutens, E., F. Rost, and G. Depre´. 1984. A nosocomial epidemic of Serratia liquefaciens urinary tract infection after cystometry. Eur. J. Clin. Microbiol. 3:316–317. 345. Severn, M. 1977. Gentamicin-resistant Serratia. Lancet 309:750. 346. Shanks, R. M. Q., et al. 2007. A Serratia marcescens OxyR homolog mediates surface attachment and biofilm formation. J. Bacteriol. 189:7262–7272. 347. Shaw, K. J., et al. 1992. Characterization of the chromosomal aac(6⬘)-Ic gene from Serratia marcescens. Antimicrob. Agents Chemother. 36:1447– 1455. 348. Sheng, W.-H., et al. 2002. Emerging fluoroquinolone-resistance for common clinically important gram-negative bacteria in Taiwan. Diagn. Microbiol. Infect. Dis. 43:141–147. 349. Shi, Z.-Y., P. Y.-F. Liu, Y.-J. Lau, Y.-H. Lin, and B.-S. Hu. 1997. Use of pulsed-field gel electrophoresis to investigate an outbreak of Serratia marcescens. J. Clin. Microbiol. 35:325–327. 350. Shimizu, K., et al. 1985. Comparison of aminoglycoside resistance patterns in Japan, Formosa, and Korea, Chile, and the United States. Antimicrob. Agents Chemother. 28:282–288. 351. Shimuta, K., et al. 2009. The hemolytic and cytolytic activities of Serratia marcescens phospholipase A (PhlA) depend on lypophospholipid production by PhlA. BMC Microbiol. 9:261.

790

MAHLEN

352. Sidjabat, H. E., et al. 2009. Interspecies spread of Klebsiella pneumoniae carbapenemase gene in a single patient. Clin. Infect. Dis. 49:1736–1738. 353. Siegman-Igra, Y., G. Inbar, and A. Campus. 1985. An ‘outbreak’ of pulmonary pseudoinfection by Serratia marcescens. J. Hosp. Infect. 6:218–220. 354. Sikka, M. K., et al. 2010. Microbiologic and clinical epidemiologic characteristics of the Chicago subset of a multistate outbreak of Serratia marcescens bacteremia. Infect. Control Hosp. Epidemiol. 31:1191–1193. 355. Silva, C. V., et al. 2003. Pseudo-outbreak of Pseudomonas aeruginosa and Serratia marcescens related to bronchoscopes. Infect. Control Hosp. Epidemiol. 24:195–197. 356. Simor, A. E., L. Ramage, L. Wilcox, S. B. Bull, and H. Bialkowska-Hobrzanska. 1988. Molecular and epidemiologic study of multiresistant Serratia marcescens infections in a spinal cord injury rehabilitation unit. Infect. Control 9:20–27. 357. Singh, B. R., Y. Singh, and A. K. Tiwari. 1997. Characterization of virulence factors of Serratia strains isolated from foods. Int. J. Food Microbiol. 34: 259–266. 358. Skerman, V. B. D., V. McGowan, and P. H. A. Sneath. 1980. Approved lists of bacterial names. Int. J. Syst. Bacteriol. 30:225–420. 359. Smith, P. J., D. S. Brookfield, D. A. Shaw, and J. Gray. 1984. An outbreak of Serratia marcescens infections in a neonatal unit. Lancet 323:151–153. 360. Sokalski, S. J., M. A. Jewell, A. C. Asmus-Shillington, J. Mulcahy, and J. Segreti. 1992. An outbreak of Serratia marcescens in 14 adult cardiac surgical patients associated with 12-lead electrocardiogram bulbs. Arch. Intern. Med. 152:841–844. 361. Soler-Palacín, P., et al. 2007. Chronic granulomatous disease in pediatric patients: 25 years of experience. Allergol. Immunopathol. (Madrid) 35: 83–89. 362. Stamm, W. E., et al. 1976. A nursery outbreak caused by Serratia marcescens—scalp-vein needles as a portal of entry. J. Pediatr. 89:96–99. 363. Stapp, C. 1940. Bacterium rubidaeum nov. sp. Zentralbl. Bakteriol. II Abt. 102:251–260. 364. Stenderup, A., O. Faergeman, and M. Ingerslev. 1966. Serratia marcescens infections in premature infants. Acta Pathol. Microbiol. Scand. 68:157–160. 365. Stephen, M., and M. K. Lalitha. 1993. An outbreak of Serratia marcescens infection among obstetric patients. Indian J. Med. Res. 97:202–205. 366. Steppberger, K., et al. 2002. Nosocomial neonatal outbreak of Serratia marcescens—analysis of pathogens by pulsed field gel electrophoresis and polymerase chain reaction. Infection 30:277–281. 367. Stock, I., S. Burak, K. J. Sherwood, T. Gru ¨ger, and B. Wiedemann. 2003. Natural antimicrobial susceptibilities of ‘unusual’ Serratia species: S. ficaria, S. fonticola, S. odorifera, S. plymuthica, and S. rubidaea. J. Antimicrob. Chemother. 51:865–885. 368. Stock, I., T. Grueger, and B. Wiedemann. 2003. Natural antibiotic susceptibility of strains of Serratia marcescens and the S. liquefaciens complex: S. liquefaciens sensu stricto, S. proteamaculans, and S. grimesii. Int. J. Antimicrob. Agents 22:35–47. 369. Su, L. H., et al. 2003. Extended epidemic of nosocomial urinary tract infections caused by Serratia marcescens. J. Clin. Microbiol. 41:4726–4732. 370. Su, J. R., et al. 2009. Epidemiologic investigation of a 2007 outbreak of Serratia marcescens bloodstream infection in Texas caused by contamination of syringes prefilled with heparin and saline. Infect. Control Hosp. Epidemiol. 30:593–595. 371. Suh, B., et al. 2010. Outbreak of meropenem-resistant Serratia marcescens comediated by chromosomal AmpC beta-lactamase overproduction and outer membrane protein loss. Antimicrob. Agents Chemother. 54:5057– 5061. 372. Sunenshine, R. H., et al. 2007. A multistate outbreak of Serratia marcescens bloodstream infection associated with contaminated intravenous magnesium sulfate from a compounding pharmacy. Clin. Infect. Dis. 45:527–533. 373. Sutherland, K. P., et al. 2010. Human sewage identified as likely source of white pox disease of the threatened Caribbean elkhorn coral, Acropora palmata. Environ. Microbiol. 12:1122–1131. 374. Takahashi, H., et al. 2004. Nosocomial Serratia marcescens outbreak in Osaka, Japan, from 1999 to 2000. Infect. Control Hosp. Epidemiol. 25:156– 161. 375. Tanaka, T., H. Takahashi, J. M. Kobayashi, T. Ohyama, and N. Okabe. 2004. A nosocomial outbreak of febrile bloodstream infection caused by heparinized-saline contaminated with Serratia marcescens, Tokyo, 2002. Jpn. J. Infect. Dis. 57:189–192. 376. Taylor, G., and P. M. Keane. 1962. Cross-infection with Serratia marcescens. J. Clin. Pathol. 15:145–147. 377. Thayer, H. H. 1966. Contraindication for use of Serratia marcescens as tracer organisms in research. J. Dent. Res. 45:853–855. 378. Thomas, F. E., R. T. Jackson, A. Melly, and R. H. Alford. 1977. Sequential hospitalwide outbreaks of resistant Serratia and Klebsiella infections. Arch. Intern. Med. 137:581–584. 379. Thomas, F. E., Jr., J. M. Leonard, and R. H. Alford. 1976. Sulfamethoxazole-trimethoprim-polymyxin therapy of serious multiply drug-resistant Serratia infections. Antimicrob. Agents Chemother. 9:201–207. 380. Thompson, S. A., E. V. Maani, A. H. Lindell, C. J. King, and J. V. McArthur. 2007. Novel tetracycline resistance determinant isolated from an en-

CLIN. MICROBIOL. REV.

381.

382.

383.

384.

385.

386.

387.

388. 389.

390.

391.

392. 393.

394.

395.

396.

397.

398.

399.

400. 401. 402. 403.

404.

405.

406. 407.

408. 409.

vironmental strain of Serratia marcescens. Appl. Environ. Microbiol. 73: 2199–2206. Thomson, N. R., M. A. Crow, S. J. McGowan, A. Cox, and G. P. Salmond. 2000. Biosynthesis of carbapenem antibiotic and prodigiosin pigment in Serratia is under quorum sensing control. Mol. Microbiol. 36:539–556. Torii, K., Y. Noda, Y. Miyazaki, and M. Ohta. 2003. An unusual outbreak of infusion-related bacteremia in a gastrointestinal disease ward. Jpn. J. Infect. Dis. 56:177–178. Traub, W. H. 1972. Continued surveillance of Serratia marcescens infections by bacteriocin typing: investigation of two outbreaks of cross-infection in an intensive care unit. Appl. Microbiol. 23:982–985. Tsakris, A., et al. 1992. Prevalence of a plasmid-mediated type II dihydrofolate reductase gene among trimethoprim-resistant urinary pathogens in Greek hospitals. J. Antimicrob. Chemother. 29:405–413. Tsakris, A., et al. 2010. In vivo acquisition of a plasmid-mediated blaKPC-2 gene among clonal isolates of Serratia marcescens. J. Clin. Microbiol. 48: 2546–2549. Turner, P. J. 2009. MYSTIC Europe 2007: activity of meropenem and other broad-spectrum agents against nosocomial isolates. Diagn. Microbiol. Infect. Dis. 63:217–222. Uduman, S. A., et al. 2002. An outbreak of Serratia marcescens infection in a special-care baby unit of a community hospital in United Arab Emirates: the importance of the air conditioner duct as a nosocomial reservoir. J. Hosp. Infect. 52:175–180. Ursua, P. R., et al. 1996. Serratia rubidaea as an invasive pathogen. J. Clin. Microbiol. 34:216–217. Vandenbroucke-Grauls, C. M., A. C. Baars, M. R. Visser, P. F. Hulstaert, and J. Verhoef. 1993. An outbreak of Serratia marcescens traced to a contaminated bronchoscope. J. Hosp. Infect. 23:263–270. van der Sar-van der Brugge, S., et al. 1999. Risk factors for acquisition of Serratia marcescens in a surgical intensive care unit. J. Hosp. Infect. 41:291– 299. van der Vorm, E. R., and C. Woldring-Zwaan. 2002. Source, carriers, and management of a Serratia marcescens outbreak on a pulmonary unit. J. Hosp. Infect. 52:263–267. Van Houdt, R., M. Givskov, and C. W. Michiels. 2007. Quorum sensing in Serratia. FEMS Microbiol. Rev. 31:407–424. van Ogtrop, M. L., D. van Zoeren-Grobben, E. M. A. Verbakel-Salomons, and C. P. A. van Boven. 1997. Serratia marcescens infections in neonatal departments: description of an outbreak and review of the literature. J. Hosp. Infect. 36:95–103. Velasco, C., et al. 2010. Smaqnr, a new chromosome-encoded quinolone resistance determinant in Serratia marcescens. J. Antimicrob. Chemother. 65:239–242. Vigeant, P., et al. 1998. An outbreak of Serratia marcescens infections related to contaminated chlorhexidine. Infect. Control Hosp. Epidemiol. 19:791–794. Villari, P., M. Crispino, A. Salvadori, and A. Scarcella. 2001. Molecular epidemiology of an outbreak of Serratia marcescens in a neonatal intensive care unit. Infect. Control Hosp. Epidemiol. 22:630–634. Villarino, M. E., W. R. Jarvis, C. O’Hara, J. Bresnahan, and N. Clark. 1989. Epidemic of Serratia marcescens bacteremia in a cardiac intensive care unit. J. Clin. Microbiol. 27:2433–2436. Voelz, A., et al. 2010. Outbreaks of Serratia marcescens in neonatal and pediatric intensive care units: clinical aspects, risk factors and management. Int. J. Hyg. Environ. Health 213:79–87. Waisman, H. A., and W. H. Stone. 1958. The presence of Serratia marcescens as the predominating organism in the intestinal tract of the newborn. Pediatrics 21:8–12. Wake, C., H. Lees, and A. B. Cull. 1986. The emergence of Serratia marcescens as a pathogen in a newborn unit. Aust. Paediatr. J. 22:323–326. Waldermann, F., O. M. Maucher, A. Buck, and E. Goerttler. 1987. Fistulous pyoderma caused by Serratia liquefaciens. Hautarzt 38:36–39. (In German.) Walther-Rasmussen, J., and N. Høiby. 2007. Class A carbapenemases. J. Antimicrob. Chemother. 60:470–482. Walther-Wenke, G., et al. 2010. Screening of platelet concentrates for bacterial contamination: spectrum of bacteria detected, proportion of transfused units, and clinical follow-up. Ann. Hematol. 89:83–91. Washington, I. I., J. A. R. J. Birk, and R. E. Ritts, Jr. 1971. Bacteriologic and epidemiologic characteristics of Enterobacter hafniae and Enterobacter liquefaciens. J. Infect. Dis. 124:379–386. Weigel, L. M., C. D. Steward, and F. C. Tenover. 1998. gyrA mutations associated with fluoroquinolone resistance in eight species of Enterobacteriaceae. Antimicrob. Agents Chemother. 42:2661–2667. Whalen, T. A. 1970. Serratia marcescens: a pathogen. Science 168:64. Wheat, R. P., A. Zuckerman, and L. A. Rantz. 1951. Infection due to chromobacteria; a report of eleven cases. AMA Arch. Intern. Med. 88:461– 466. Whitby, J. L., J. N. Blair, and A. Rampling. 1972. Cross-infection with Serratia marcescens in an intensive-therapy unit. Lancet 300:127–129. Wilhelmi, I., et al. 1987. Epidemic outbreak of Serratia marcescens infection in a cardiac surgery unit. J. Clin. Microbiol. 25:1298–1300.

VOL. 24, 2011 410. Williamson, N. R., P. C. Fineran, F. J. Leeper, and G. P. C. Salmond. 2006. The biosynthesis and regulation of bacterial prodiginines. Nat. Rev. 4:887– 899. 411. Winslow, C.-E. A., et al. 1920. The families and genera of the bacteria. Final report of the Committee of the Society of American Bacteriologists on characterization and classification of bacterial types. J. Bacteriol. 5:191– 229. 412. Woodfield, D. G. 1991. Transfusion acquired Serratia liquefaciens septicaemia. N. Z. Med. J. 104:141. 413. Woodward, H. M. M., and K. B. Clarke. 1913. A case of infection in man by the Bacterium prodigiosum. Lancet 181:314–315. 414. Wu, L. T., et al. 2004. Survey of CTX-M-3 extended-spectrum ␤-lactamase (ESBL) among cefotaxime-resistant Serratia marcescens at a medical center in middle Taiwan. Diagn. Microbiol. Infect. Dis. 49:125–129. 415. Yamamoto, T., A. Ariyoshi, and K. Amako. 1985. Fimbria-mediated adherence of Serratia marcescens strain US5 to human urinary bladder surface. Microbiol. Immunol. 29:677–681. 416. Yamazaki, G., et al. 2006. Effect of salt stress on pigment production of Serratia rubidaea N-1: a potential indicator strain for screening quorum sensing inhibitors from marine microbes. J. Gen. Appl. Microbiol. 52:113– 117. 417. Yang, Y. J., P. J. Wu, and D. M. Livermore. 1990. Biochemical characterization of a beta-lactamase that hydrolyzes penems and carbapenems from two Serratia marcescens isolates. Antimicrob. Agents Chemother. 34:755– 758. 418. Yoon, H. J., et al. 2005. Outbreaks of Serratia marcescens bacteriuria in a neurosurgical intensive care unit of a tertiary care teaching hospital: a clinical, epidemiologic, and laboratory perspective. Am. J. Infect. Control 33:595–601.

Steven D. Mahlen is the Medical Director of Microbiology in the Department of Pathology and Area Laboratory Services at Madigan Healthcare System in Tacoma, WA. Dr. Mahlen received his B.S. and M.S. in microbiology from South Dakota State University and his Ph.D. in medical microbiology from Creighton University, has completed a clinical microbiology fellowship at the University of Washington, and is a Diplomate of the American Board of Medical Microbiology. He is a lieutenant colonel in the U.S. Army and has been on active duty since 1993. During this time, Dr. Mahlen has directed two different Army clinical microbiology laboratories in the United States, has performed a variety of assignments, and has been the Chief of the Laboratory in combat support hospitals on two different deployments: 1995–1996, in support of the United Nations Mission in Haiti, and 2006–2007, in support of Operation Iraqi Freedom.

SERRATIA INFECTIONS

791

419. Yu, V. L. 1979. Serratia marcescens: historical perspective and clinical review. N. Engl. J. Med. 300:887–893. 420. Yu, W.-L., et al. 2008. Institutional spread of clonally related Serratia marcescens isolates with a novel AmpC cephalosporinase (S4): a 4-year experience in Taiwan. Diagn. Microbiol. Infect. Dis. 61:460–467. 421. Yu, W. L., L. T. Wu, M. A. Pfaller, P. L. Winokur, and R. N. Jones. 2003. Confirmation of extended-spectrum beta-lactamase-producing Serratia marcescens: preliminary report from Taiwan. Diagn. Microbiol. Infect. Dis. 45:221–224. 422. Yum, J. H., D. Yong, K. Lee, H.-S. Kim, and Y. Chong. 2002. A new integron carrying VIM-2 metallo-␤-lactamase gene cassette in a Serratia marcescens isolate. Diagn. Microbiol. Infect. Dis. 42:217–219. 423. Zaidi, M., J. Sifuentes, M. Bobadilla, D. Moncada, and S. Ponce de Leo´n. 1989. Epidemic of Serratia marcescens bacteremia and meningitis in a neonatal unit in Mexico City. Infect. Control Hosp. Epidemiol. 10:14–20. 424. Zbinden, R., and R. Blass. 1988. Serratia plymuthica osteomyelitis following a motorcycle accident. J. Clin. Microbiol. 26:1409–1410. 425. Zhang, C.-X., et al. 2009. Serratia nematodiphila sp. nov., associated symbiotically with the entomopathogenic nematode Heterorhabditidoides chongmingensis (Rhabditida: Rhabditidae). Int. J. Syst. Evol. Microbiol. 59:1603–1608. 426. Zhang, R., H. W. Zhou, J. C. Cai, and G. X. Chen. 2007. Plasmid-mediated carbapenem-hydrolysing beta-lactamase KPC-2 in carbapenem-resistant Serratia marcescens isolates from Hangzhou, China. J. Antimicrob. Chemother. 59:574–576. 427. Zobell, C. E., and H. C. Upham. 1944. A list of marine bacteria, including descriptions of sixty new species. Bull. Scripps Inst. Oceanogr. Univ. Calif. Tech. Ser. 5:239–292.

CLINICAL MICROBIOLOGY REVIEWS, Oct. 2011, p. 792–805 0893-8512/11/$12.00 doi:10.1128/CMR.00014-11 Copyright © 2011, American Society for Microbiology. All Rights Reserved.

Vol. 24, No. 4

Immunodiagnosis of Tuberculosis: a Dynamic View of Biomarker Discovery Shajo Kunnath-Velayudhan and Maria Laura Gennaro* The Public Health Research Institute, New Jersey Medical School, 225 Warren Street, Newark, New Jersey 07103 INTRODUCTION .......................................................................................................................................................792 THE GRANULOMATOUS LESIONS .....................................................................................................................793 The Tuberculous Granuloma ................................................................................................................................793 Priming in the Granuloma ....................................................................................................................................794 Bacterial Phenotypes and Antigen Expression ...................................................................................................795 Local and Systemic Immune Responses..............................................................................................................795 IMMUNE MARKERS ................................................................................................................................................796 Mediators of Antigen-Specific Immune Responses............................................................................................796 Cellular Immune Responses and Infection State...............................................................................................797 Cellular responses in latent infection and active TB ....................................................................................797 Cellular responses and antigen expression.....................................................................................................798 Cellular responses and disease progression ...................................................................................................798 Humoral Immune Responses and Infection State .............................................................................................798 Antibody responses and bacterial metabolic status.......................................................................................798 Antibody responses and bacterial burden .......................................................................................................799 Antibody responses and disease progression..................................................................................................799 Diversity of antibody responses ........................................................................................................................799 NEW PRINCIPLES FOR BIOMARKER DISCOVERY........................................................................................800 A Paradigm Shift ....................................................................................................................................................800 Systems Immunology ..............................................................................................................................................800 CONCLUSIONS .........................................................................................................................................................801 ACKNOWLEDGMENTS ...........................................................................................................................................801 REFERENCES ............................................................................................................................................................801 Diagnosing TB is no simple matter. Infection with M. tuberculosis has been commonly regarded as having a binary clinical outcome. One is LTBI, which is characterized by a positive tuberculin skin test (TST)/IGRA, the absence of symptoms, and a normal chest X-ray. The other outcome is active TB, which is typically defined by the detection of tubercle bacilli or bacillary products in pathological specimens, usually sputum. However, it is becoming increasingly clear that the clinical spectrum of M. tuberculosis infection is more complex than previously appreciated. The definition of LTBI includes multiple conditions, as is best recognized in nonhuman primate models (6, 162). In humans, forms of LTBI can be differentiated on the basis of risk of reactivation. For example, in immunocompetent individuals, the annual risk of developing active TB is 1.5% in the first 2 years after infection and 0.1% thereafter (96). Asymptomatic infection with a history of past TB carries a greater risk of reactivation than LTBI alone, particularly when chest X-ray findings are abnormal (104). Active pulmonary TB also presents with a spectrum of clinical manifestations, which are usually associated with increasing bacillary burden. Since active TB is diagnosed with bacteriological assays, a low bacillary burden often leads to a missed diagnosis. It emerges from the above considerations that M. tuberculosis infection results in a continuum of ill-defined, sometimes overlapping, clinical manifestations (6, 162) (Fig. 1). Since treatment decisions are based on particular criteria (for example, treatment of LTBI is warranted only when the risk of reactivation is high [3]), recognizing the spectrum of

INTRODUCTION The importance of diagnostic research on tuberculosis (TB) cannot be overstated. One-third of the world population carries an asymptomatic infection with Mycobacterium tuberculosis, which results in eight million new cases of TB and two million deaths every year. Identifying and treating those who progress to disease and can transmit infection to contacts are crucial to successful control. Our current diagnostic toolbox is inadequate to achieve these goals. For example, none of the tests for active TB is sufficiently accurate, timely, and appropriate for low-income and low-technology settings, where most TB cases are found (reviewed in references 44, 87, 88, 91, 143, 144, 145, and 162). The recently developed gamma interferon (IFN-␥) release assays (IGRAs) diagnose latent M. tuberculosis infection (LTBI) more accurately than the century-old tuberculin skin test (reviewed in references 36, 77, 109, and 110), but they fail to facilitate decisions concerning targeted LTBI treatment (97, 107, 108). Indeed, it is recognized that the millennium development goals set by the United Nations in the fight against TB, i.e., cutting in half the global prevalence and death rate by 2015 (42), cannot be reached without the development of new diagnostic tools (http://www.stoptb.org/globalplan).

* Corresponding author. Mailing address: The Public Health Research Institute, New Jersey Medical School, 225 Warren Street, Newark, NJ 07103. Phone: (973) 854-3210. Fax: (973) 854-3101. E-mail: [email protected]. 792

VOL. 24, 2011

IMMUNODIAGNOSIS OF TUBERCULOSIS

793

FIG. 1. Clinical states of M. tuberculosis infection. This schematic is adapted from the classification of TB by the American Thoracic Society (ATS) (2). ATS class numbers are also indicated, as applicable. Infected individuals are divided into asymptomatic and symptomatic. (i) The asymptomatic group is further divided into subgroups; color codes indicate the relative risk of progression to active disease in each subgroup (green, low; yellow, high). Past TB (inactive TB; class 4) indicates either a history of a previous episode(s) of active TB or abnormal stable radiographic findings and no bacteriological and/or radiographic evidence of current disease. LTBI (class 1) indicates a positive TST/IGRA and no clinical, bacteriological, or radiographic evidence of active disease. LTBI is further divided into incident/recent (⬍2 years after infection) or prevalent/remote (⬎2 years postinfection). The preclinical TB/incipient TB group includes asymptomatic individuals found to have developed active disease when examined at later (short-term) times. (ii) The symptomatic group is also further divided into subgroups; here, color codes indicate bacillary load (orange, low; red, high). Clinical TB indicates symptoms and/or radiographic findings suggestive of active TB but no bacteriological evidence of disease. Culture-confirmed TB (class 3) indicates bacteriological evidence of active TB. These patients are further subdivided into smear-negative and smear-positive groups based on sputum smear microscopy (It is noted that the extent of radiographic lung involvement, such as cavitary and noncavitary disease, is often also used to classify patients.).

asymptomatic and symptomatic stages of M. tuberculosis infection is critical for implementing more effective TB control policies. Immunological biomarkers should best distinguish the stages of M. tuberculosis infection from one another. Immunological events are at the core of TB pathogenesis, since they are responsible for both tissue damage and protection (41). Thus, various phases and outcomes of M. tuberculosis infection should be associated with particular immunological events. Moreover, assessing immune responses circumvents the need to detect tubercle bacilli or their products, both of which are currently inaccessible during most of the asymptomatic infection and even during early symptomatic stages. Despite these considerations, no immunodiagnostic test exists that can accurately diagnose active TB, distinguish LTBI from active TB, or tell apart asymptomatic forms of infection that are associated with a high risk of disease progression (the shortcomings of TB immunodiagnostics have been extensively reviewed [36, 38, 134, 142, 143]). It is time to translate the complexities of the clinical spectrum of M. tuberculosis infection into new paradigms for TB immunodiagnosis. The present review explores parallels between the clinical and pathological development of M. tuberculosis infection and the evolution of immune responses. Previous considerations of the immunological spectrum in TB have usually been limited to clinical manifestations of disease (see, for example, references 29 and 81), typically in search of a parallel with leprosy (100, 119). These have been guided primarily by the Th1/Th2 paradigm, which is now seen as an oversimplification of opposite immunophenotypes (61). Here we examine the evolution of histopathological events, immune mediators, and cell types in peripheral blood and their relationship to the physiology of the tubercle bacillus (Fig. 2) (we do not review the effects of coinfection with HIV, which relate primarily to loss of immune control during latent M. tuberculosis infection). We then discuss how recognizing the immunological spectrum of M. tuberculosis infection impacts the development of new immunodiagnostics. The present article emphasizes the need for

“combinatorial” systems approaches to biomarker selection rather than the diagnostic characteristics of individual biomarkers. For excellent meta-analyses of immunological biomarkers and their reported accuracy in the diagnosis of LTBI and/or active TB, the reader is referred to recent reviews (162, 163). THE GRANULOMATOUS LESIONS Tubercle bacilli interact with immune cells and various host cell types interact with each other to form the granuloma, a dynamic structure that is the histopathological hallmark of M. tuberculosis infection. The granuloma acts as a site of immune cell priming. The systemic immune response should reflect changes in the local immune compartments due to the recirculation of immune cells and the release of soluble mediators that reach the periphery. The Tuberculous Granuloma The tuberculous granuloma is a dynamic multicellular aggregate. Macrophages predominate in the inner cellular layer as multinucleated giant cells, epithelioid cells, or foamy cells (127). T lymphocytes expressing ␣␤ T-cell receptors (TCR) are also present. CD4⫹ T cells are localized in both inner and outer cellular layers, while CD8⫹ T cells are found mostly in the outer cellular layer. The outer cellular layer also contains B cell-rich aggregates, which include bacillus-laden antigenpresenting cells (APCs) and T lymphocytes. Granulomas also contain atypical lymphocytes such as ␥␦ T cells, which do not require major histocompatibility complex (MHC) class I or II molecules for antigen recognition, and regulatory T cells with immunosuppressive properties (12, 58). The typical caseous granuloma includes a central necrotic core. Some bacilli are found in the necrotic core, but most are located at the interface between the necrotic core and the macrophage-rich, inner cellular layer (126, 156, 158). As disease progresses, granulomas tend to be less organized and may become cavitary (157).

794

KUNNATH-VELAYUDHAN AND GENNARO

CLIN. MICROBIOL. REV.

FIG. 2. Schematic representation of bacteriological, histopathological, and immunological changes during M. tuberculosis infection. The background color reflects the clinical spectrum of infection, progressing from asymptomatic (yellow) to symptomatic (red). The changes in the granulomatous lesions are shown relative to the number of lesions (vertical axis) and the quality of the granuloma (color composition). In each granuloma icon, the gray, granular area represents cells, while the solid orange color represents caseum. For tubercle bacilli, bacterial numbers (vertical axis) and phenotype (color composition of the bacilli) are depicted. In each icon, the color indicates the growth phase (green, growing bacilli; red, nongrowing bacilli). For antibody levels, the icon, which represents antibody responses to the entire proteome, is roughly divided into a nonreactive, dominant area of the proteome (outer, colorless) an and inner, reactive area (red gradient). The gradient of red indicates rarely reactive (orange) and commonly reactive (dark red) proteins. The height of the reactive proteome area (vertical axis) represents the frequency of reactive TB sera. The early, transient peak shown in the antibody curve is derived from monkey and human data. A similar course of the bacillary curve is inferred from the likelihood that tubercle bacilli multiply before immunity is expressed and bacillary growth is controlled. For cytokine levels, three hypothetical patterns are shown, with levels (vertical axis) decreasing (purple) or increasing (brown) with disease progression or being detected only during active disease (green). Each pattern may be characteristic of one or more cytokines.

When it occurs, cavitation connects the caseous center with the bronchial tree. Loss of organization in the granuloma alters the interactions between immune cells (75) and correlates with tissue destruction, as indicated in active TB by increased serum levels of matrix metalloproteinase-9 (65), a protein that degrades extracellular matrix (115). Granuloma progression is associated with increased bacterial numbers, particularly at the luminal surface of the cavity. This may be favored by the local selective reduction of T cells, which diminishes T-cell–macrophage interactions (7, 75, 157). Disorganization of advanced granulomas may also facilitate bacterial dissemination to other areas of the lung and to other organs, with formation of new granulomas (157). There are many variants of the tuberculous granuloma (86), which differ in cellular composition and structure. Some granulomas are rich in granulocytes (86). Others vary with regard to T-cell composition. For example, it was reported that while most granulomas stained positive for CD4⫹ T cells, only half were positive for CD8⫹ T cells (46). Consequently, each granuloma may exhibit distinct profiles of cytokine production (45, 46). When a necrotic center is present, its characteristics may also vary (e.g., central necrosis may be suppurative). Healing of granulomas is associated with fibrotic changes, mineralization (15, 86), and presumably the presence of few, if any, tubercle bacilli. Differences between granulomas are also seen in the pattern of distribution in the lung, which may be focal, multifocal, coalescing, or invasive. While the architecture of the

granuloma typically changes with the evolution of infection, different types of granulomatous lesions are also found in the same individual, either with active disease (47) or with latent infection (6). Studies on experimentally infected macaques have greatly contributed to understanding the evolution of the granuloma structure with disease progression (15, 86). In these animals, latent infection is usually associated with one or few localized granulomas, with or without minimal involvement of thoracic lymph nodes. Active disease is characterized by large numbers of highly disseminated granulomas, which tend to invade nearby vessels and airways. Granulomas in animals with latent infection differ from those found in animals with active disease with regard to structure and immune cell composition: active disease is associated with higher numbers of caseous granulomas, more numerous CD4⫹ and CD8⫹ T cells (up to 100-fold), and higher proportions of T cells expressing the chemokine receptors CCR5 and CXCR3 (86). Differences are also seen with antigen-specific immune responses, as the number of cells producing IFN-␥ in response to the M. tuberculosis secreted proteins ESAT-6 and CFP-10 is higher in the lungs of monkeys with active disease than in those with latent infection (86). Priming in the Granuloma The granuloma is a site of immunological priming, which occurs at the interface between the macrophage-rich inner

VOL. 24, 2011

layer and the surrounding T-cell-rich outer layer (126, 156, 158). Since T cells in this layer are predominantly CD4⫹ cells, they are more likely to interact with APCs than CD8⫹ T cells, which are found mostly in the outer cellular layer. Additional priming sites for B cells and T cells include the follicle-like structures found in the outer cellular layer, which are rich in B cells, CD4⫹ and CD8⫹ T cells, and infected APCs. In these follicles, B cells are found at different states of differentiation (naïve cells, memory cells, and antibody-secreting plasma cells). An additional route of priming may exist in caseous granulomas, where mycobacterial antigens presumably released from dead bacilli in the necrotic center (45, 66) reach the surrounding APC-rich areas. Multiple factors at the site of priming affect the immune response. The relative local abundance of a particular cell type targeted for priming is important for the strength and persistence of the immune responses induced. The nature of the bacterial antigen is also significant. For example, B-cell priming is favored in areas containing extracellular bacteria, since B cells react preferentially with particulate antigens. Additionally, the intracellular location of bacteria has a role in the endogenous processing of antigens. For the majority of intracellular mycobacteria, which are found in the phagosome, antigen presentation is directed to MHC class II molecules (32). Tubercle bacilli have also been found in the cytoplasm of phagocytes (93, 159), a location that triggers presentation by MHC class I molecules. Although it is controversial, a possibility also exists that bacterial proteins secreted by the bacteria in the phagosome enter the surrounding host cell cytoplasm (21, 150). Priming of CD8⫹ T cells may also involve blebs, which are generated by apoptosis of infected macrophages (130), or exosomes, which are membrane-bound vesicles secreted by infected macrophages (8, 10, 51). Yet another factor affecting priming and immune responses is the local concentration of bacterial antigen. This may vary not only in relation to the absolute bacterial load but also in relation to the relative bacterial antigen expression patterns that reflect microenvironmental variations between granulomas or even between different areas of the same granuloma (45, 47, 116). The relationship between bacterial growth phenotypes and bacterial antigen expression is discussed below. Bacterial Phenotypes and Antigen Expression That tubercle bacilli multiply during active disease and remain dormant during LTBI is an oversimplified scenario, since several lines of evidence suggest that multiple bacillary phenotypes are concurrently found during infection. In the case of active TB, the need for prolonged chemotherapy is partially attributable to the phenotypic drug resistance associated with the presence of nonreplicating mycobacteria (24). Indeed, nonreplicating tubercle bacilli most likely exist in granulomatous lesions irrespective of stage and structural integrity, since prolonged anti-TB chemotherapy is required even in HIV-coinfected individuals, where granulomas tend to be less organized (24). Another line of evidence for mixed bacillary phenotypes in active TB derives from microbiological studies of sputum. Up to 85% of bacilli found in sputum samples from active TB patients can be laden with neutral lipids, which are a trait of nonreplicating M. tuberculosis (49). Moreover, sputum samples

IMMUNODIAGNOSIS OF TUBERCULOSIS

795

from most TB patients contain predominantly resuscitationpromoting factor (RFP)-dependent bacteria (101). These bacteria, which are not detected by standard CFU enumeration, require treatment with RFP proteins for growth or for regrowth following dormancy (101). RFP-dependent bacilli are tolerant to rifampin and are eliminated by antibiotic treatment more slowly than colony-forming bacilli (101). Mixed bacterial phenotypes are also likely to be present during latent infection. For example, in vitro culturing of granulomatous tissue from autoptic lungs of individuals with LTBI who died from TB-unrelated pathology has yielded mycobacteria exhibiting different rates of regrowth (43). Moreover, the ability of isoniazid, a drug effective only against multiplying bacteria, to reduce the risk of disease reactivation in LTBI (23) indirectly supports the presence of multiplying bacilli during latent infection. The possibility of gathering direct evidence on the physiological state of tubercle bacilli during latent infection has been hampered primarily by the failure to determine their location. Staining of acid-fast bacilli (AFB) may not be the detection method of choice, since nonreplicating mycobacteria exhibit reduced acid-fastness due to cell wall remodeling (28). Alternative methods, such as in situ PCR methods, have detected mycobacterial DNA in alveolar and interstitial macrophages present in “normal-appearing” lung tissue (62), suggesting locations of tubercle bacilli other than the classical granuloma. Due to the difficulties described above, whether the entire bacterial population is not growing during latent infection or a growing subset of bacilli exists remains a point of debate (43). The bacillary growth state is likely to be important for stimulation of the immune response because actively growing and dormant tubercle bacilli probably express different antigen sets. These two types of bacilli differ in major metabolic pathways, protein secretory pathways, and cell wall composition (28, 30, 137, 152, 166). Moreover, protein secretion increases with bacillary growth rate (4), and secreted proteins are favored antigen targets (26). Murine studies have shown that expression of immunodominant antigens varies with bacillary growth phase (in mice, tubercle bacilli grow exponentially in the acute phase of mouse lung infection and then stop growing during chronic infection [135]). For example, during chronic infection, expression of the gene encoding the antigen 85 (Ag85) complex, which is involved in the biosynthesis of mycolic acids (9), is downregulated (136), while expression of hspX (encoding ␣-crystallin), which is part of the so-called dormancy (devR/dosR) regulon (160), is induced (135). This dichotomy fits with mycolic acid synthesis being a cellular activity associated with bacterial multiplication and expression of the devR/dosR regulon being associated with M. tuberculosis growth arrest (120, 135). Similar inferences were made from human studies in which differences in bacterial gene expression patterns were seen between tubercle bacilli grown in vitro and those found in sputa of TB patients (49) or in human tuberculous lung tissue (116). Local and Systemic Immune Responses While the study of the infection site reveals interactions between the pathogen and immune cells (reviewed in references 72 and 132), it is the characterization of the peripheral

796

KUNNATH-VELAYUDHAN AND GENNARO

immune response that should help identify stage-specific immune markers that are suitable for diagnostic development. Thus, it is imperative to understand how well the peripheral response reflects events at the infection site. Little doubt exists that immune responses expressed at the infection site and in the periphery differ quantitatively and qualitatively. Immune cells are localized at the site of infection due to clonal expansion and migration from the peripheral circulation and from lymphoid organs after priming (35, 146, 155). As a result, the number of antigen-specific lymphocytes at the infection site is, on average, 10 times greater than that in peripheral blood (70, 71, 103). Qualitative differences may also occur. For example, CD4⫹/CD8⫹ T-cell ratios have been found to be higher in the bronchoalveolar lavage fluid than in peripheral blood during active disease (154). Moreover, a broader repertoire of epitopes recognized by T cells was found in the pleural fluid than in peripheral blood (168). On the other end, communication between the infection site and the periphery undoubtedly exists, since lymphocytes circulate between lymphoid and nonlymphoid tissues and soluble immune mediators released by these cells may reach the periphery. Immune cells in the granuloma presumably enter the bloodstream through the blood vessels found at the granulomatous site. Indeed granulomas are highly vascularized, probably due to the vascular endothelial growth factor secreted by activated macrophages at the site of infection (126). In addition, lymph nodes draining pulmonary lesions are connected to the systemic circulation. These communication lines make it possible to follow the events occurring at the site of infection by sampling peripheral blood, as demonstrated by primate studies. Indeed, the frequency of antigen-specific, IFN-␥-producing cells found in granulomas and peripheral blood of macaques is greater with active disease than with latent infection (86). IMMUNE MARKERS Many studies on T- and B-cell responses during infection with M. tuberculosis have been conducted. T lymphocytes are the effectors of the protective immune response. Thus, these cells have been extensively investigated in terms of expression of antigen-specific TCR, functional markers, and cytokine production in association with controlled infection (latent infection) versus loss of immune control (active TB), most typically in relation to vaccine research. M. tuberculosis antigens that evoke T-cell responses have also been studied, with particular attention to T-cell antigens produced by virulent tubercle bacilli but not by other, nonpathogenic mycobacterial species. An important result of that work has been the development of novel immunodiagnostics for LTBI (114, 117). The next diagnostic challenge is to distinguish between stages of M. tuberculosis infection by identification of stage-specific cellular immune markers. Expression of these markers may reflect (i) changes in antigen burden and (ii) acquisition by T cells of effector functions that are causally associated with infection outcome. In the field of antibody responses, which do not confer protection, most research has been on identifying serodominant antigens for diagnosis of active TB. Serological studies have shown pronounced variability in antibody profiles between TB patients. This has made it difficult to utilize antibodies as accurate biomarkers of active TB. Nevertheless, an-

CLIN. MICROBIOL. REV.

tibody levels do reflect bacillary burden and may track relative changes in the burden of particular bacterial antigens during the course of infection. The dynamic characteristics of the antibody response have been most clearly seen with the serological interrogation of the entire M. tuberculosis proteome. The challenge in this field is how to utilize the dynamic properties of the antibody response for diagnostics. In this section, we first review the key mediators of the immune response. We then present work describing the immunological differences between active TB and LTBI and showing how immune responses change with the evolution of infection. We discuss the two arms of immunity separately, since research on cellular and humoral immune markers has been typically conducted with different goals and different study designs (only a few studies have investigated humoral and T-cell responses concurrently; see, for example, references 89 and 92). Mediators of Antigen-Specific Immune Responses As shown in animal models, CD4⫹ T cells are initially primed in the draining lymph nodes, where the bacilli are transported by dendritic cells (169). In a mouse model, priming/activation occurs about 1 week following infection; activated cells reach the lung in 2 to 4 weeks postinfection (169). The critical role of CD4⫹ T cells in controlling M. tuberculosis infection has been shown by murine studies (105) and by the dramatic increase of the incidence of active TB associated with HIV coinfection (48). CD8⫹ T cells have the same kinetics of appearance in tissues as CD4⫹ T cells in mice. CD4⫹ and CD8⫹, MHC class Ia-restricted T cells expressing ␣␤ TCR recognize protein antigens (M. tuberculosis proteins recognized by these cell types have been listed [106]). In contrast, lipids and glycolipids are presented in the context of MHC class Ib (CD1d) molecules. Unconventional CD8⫹ T cells, which carry a ␥␦ T-cell receptor, lack the fine antigen specificity of ␣␤ TCR. These cells respond to a wide variety of pathogen-derived antigens, such as lipids, phospho- and lipoproteins, and even nonpeptide phosphorylated oligonucleotides. Many cellular immune responses are mediated by cytokines secreted by the immune cells. Cytokines are typically classified into proinflammatory Th1 cytokines and anti-inflammatory Th2 cytokines. Their relative levels determine the outcome of some mycobacterial infections, such as leprosy. In the case of TB, the association between Th1/Th2 balance and infection outcome has been actively investigated, but the results remain inconclusive (94). Many Th1 cytokines are critical for controlling infection with M. tuberculosis (reviewed in references 25, 48, and 105). One is IFN-␥, as shown by studies with individuals defective in genes for IFN-␥ or its receptor and in animal models. Another is interleukin-12 (IL-12), which stimulates IFN-␥ production. A lack of this cytokine increases susceptibility to mycobacterial infections in humans. Tumor necrosis factor alpha (TNF-␣), a proinflammatory cytokine, is also required for control of the infection, as shown by the increased incidence of TB reactivation in individuals receiving antiTNF-␣ antibody treatment for unrelated disease. Other examples of proinflammatory responses are the production of IL-2, which is involved in the clonal expansion of antigen-specific T cells (125), and of the chemokine CXCL10 (IP-10), which is important in trafficking monocytes and Th1 cells to the site of

VOL. 24, 2011

inflammation. Examples of Th2 cytokines induced by mycobacterial infection include IL-4, which has an anti-inflammatory role, and IL-10 and transforming growth factor ␤ (TGF␤), which suppress T-cell responses (reviewed in references 48 and 122). Recently, a new Th cell population (Th17) has been recognized, which produces IL-17, IL17F, IL-21, and IL-22 (153). In mice, most of the IL-17 response is generated by ␥␦ T cells (25). It appears that IL-17 and Th17 cells mediate immune pathology and may have a detrimental effect in TB (153). B lymphocytes are central to the humoral adaptive immunity. They function as APCs and may have immune regulatory roles in TB (90). The production of antibody has been investigated almost exclusively in the context of biomarker research (reviewed in references 1, 112, and 142), since a protective role against TB for antibodies is generally dismissed, with few exceptions (see, for example, reference 151). Among antibody isotypes, IgG molecules have been mainly investigated, owing to their specificity for antigen. This isotype, which is T-cell dependent, is a predominant component of secondary immune responses. The relative production of the various subclasses of IgG antibodies is influenced by the presence of particular cytokines and B-cell activators (32, 67, 140). For example, the presence of IFN-␥ favors IgG2, IL-4 and IL-13 favor IgG4, while IL-10 favors IgG1 and IgG3. In addition, subclasses depend on the biochemical nature of the antigen (32, 140). For example, most antibodies against protein antigens are of the IgG1 and IgG3 subclasses. Cellular Immune Responses and Infection State The investigation of antigen-specific, T-cell-mediated immune responses relative to infection state has followed at least two approaches. By far dominant, one has been the use of ex vivo stimulation of immune cells obtained from distinct study populations (active TB patients, LTBI cases, household contacts, and uninfected controls) with one or few immunodominant M. tuberculosis antigens (for example, ESAT-6, the 19kDa lipoprotein, and Ag85B). These studies have investigated multiple characteristics of the cells responding to antigen stimulation, such as cell surface markers and production of cytokines and chemokines, to characterize immune function in relation to infection state. This methodology can be characterized as “one/few antigens, many read-outs.” In a second approach, one or a few read-outs have been utilized (e.g., the secretion of one or a few cytokines) to assess the response of immune cells to various antigens that, for example, are presumed to be differentially expressed at various stages of infection (“many antigens, one/few read-outs”). Together, the two approaches have shown that different stages of M. tuberculosis infection are associated with quantitative and/or qualitative differences in immune cell types and released immune mediators. These are reviewed below. Cellular responses in latent infection and active TB. Cellular responses to M. tuberculosis antigens have been investigated mostly by flow cytometry analysis. One area of research has been the characterization of memory phenotypes by measuring production of IFN-␥ and IL-2, given that effector cells secrete predominantly IFN-␥, effector-memory cells secrete both IFN-␥ and IL-2, and central memory cells are known to secrete

IMMUNODIAGNOSIS OF TUBERCULOSIS

797

only IL-2 (99). In active TB, most antigen-specific T cells are effector cells, while central memory cells predominate in LTBI (16, 129). Antibiotic treatment causes a relative decrease of cells expressing effector phenotypes and a concurrent increase of memory cells in treated TB patients (16, 99). These results suggest that the relative frequencies of memory cells and effector cells are associated with changes in antigen burden, as described for viral infections (111). Another line of investigation has characterized multifunctional T cells (34, 133). Multifunctional T cells secreting IFN-␥, TNF-␣, and IL-2 are more frequent in TB patients (85 to 90%) than in LTBI cases (10 to 15%) (14, 147). Their frequency decreases during antituberculosis treatment (14, 171). Thus, also the abundance of multifunctional T cells may track increased bacterial load associated with development of active disease (14). In another study, the frequency of T cells producing only TNF-␣ was sufficient to distinguish active TB from LTBI (59). Together, these studies indicate that the responses of circulating antigen-specific T cells differ between active TB and latent infection. Even though flow cytometry analysis is highly informative, methods detecting soluble immune mediators as solid-phase antigen are more suitable for diagnostic development. The cytokine most investigated in TB research has been IFN-␥. The success of IGRAs in diagnosing LTBI has encouraged new effort to determine whether the presence of active TB correlates with increased levels of IFN-␥ relative to those detected in LTBI. Studies with experimentally infected animals support this possibility (5). Indeed, many studies conducted with IGRAs have shown higher IFN-␥ responses in active TB patients than in LTBI cases (17, 73, 74, 164). Discordant results, however, have also been obtained (113). The disease-associated increase seems to be more evident with the enzyme-linked immunospot (ELISPOT) assay, an assay enumerating IFN-␥producing cells, than with enzyme-linked immunosorbent assay (ELISA), which measures the levels of IFN-␥ secreted by the cultured cells (17, 18, 39, 40, 79, 80, 118). Thus, the two assays may have different sensitivities. Despite these encouraging results, it is difficult to distinguish active TB from latent infection on the basis of IFN-␥ levels alone (97), due to individual variability of IFN-␥ levels and the resulting overlap between the two states. Moreover, the finding that IGRA-negative contacts of TB cases can progress to active TB in less than 2 years after exposure (170) suggests perhaps that either the antigens used or the cytokine measured in the IGRAs may not cover the entire population. In addition, the sensitivity of IGRAs appears to be lower in active TB than in LTBI (see, for example, references 124 and 125), presumably due to immunosuppression accompanying active disease or to sequestration of IFN-␥-producing cells at the infection site (78). Thus, IGRAs cannot be used to rule out active TB (95). It has been suggested that the use of particular ESAT-6-derived peptides in lieu of full-length protein may enhance discrimination between active disease and latent infection (53, 56). Together with the observation that responses to ex vivo stimulation with purified protein derivative (PPD) do not differ between active TB and LTBI (64), the results obtained with ESAT-6 peptides suggest that the choice of epitope is a critical factor in the evaluation of biomarker levels in response-to-antigen assays. Measurement of additional cytokines secreted by peripheral

798

KUNNATH-VELAYUDHAN AND GENNARO

blood mononuclear cells (PBMCs) in response to M. tuberculosis antigens may also help distinguish active TB from LTBI. In one study, levels of IL-2 measured after prolonged incubation (72 h) of PBMCs with the antigens used in IGRAs were higher in LTBI than in active disease (11). The diagnostic potential of IP-10 (125) also has been evaluated, with mixed results (54, 149, 167). Since secretion of IP-10 is less affected than that of IFN-␥ by immunosuppression (55) and is age independent (85), IP-10 has been proposed as an adjunct marker of LTBI diagnosis in pediatric and HIV-coinfected populations (85, 124). In other studies, the levels of epidermal growth factor (EGF), sCD40L, vascular endothelial growth factor (VEGF), TGF-␣, and IL-1␣ released by antigen-stimulated PBMCs in a commercial IGRA distinguished TB patients from household contacts more accurately than any single marker alone (19). Release of multiple cytokines in response to bacterial antigens other than those used for the commercial IGRAs may also increase diagnostic accuracy (see, for example, references 20 and 148). Indeed, cellular responses vary with the antigen used to stimulate the cells ex vivo (60, 123). While of limited scope, these studies show that multiple host markers can help discriminate between infection states. Cellular responses and antigen expression. Few studies have investigated the correlation between cellular responses and bacterial antigen expression. Murine studies have shown that in the peripheral blood of infected animals, the number of CD4⫹ T cells reacting to ESAT-6 is higher than that of Ag85Bspecific CD4⫹ T cells; this correlates with greater abundance of the ESAT-6 transcript relative to the Ag85B transcript in lung RNA (121). The evidence is indirect in humans, where only few studies have investigated this correlation. Immune recognition of bacterial antigens associated with the nonreplicating bacterial state (“dormancy” antigens) is stronger in latent infection than in active disease, presumably correlating with higher expression of these antigens during latent infection. For example, individuals having latent infection express stronger responses to antigens encoded by the dosR regulon than active TB patients (83). A protein encoded by the dosR regulon (Rv3407) has been proposed as a latent infection marker (131). Furthermore, the cellular immune response to Rv2628, another product of the dosR regulon, was associated with cured TB and remote infection (52). Moreover, cellular responses to ␣-crystallin, which is encoded by another dosRregulated gene, are stronger in LTBI than in active TB (33). Additionally, IFN-␥ production in response to Rv2659 and Rv2660, which are two proteins encoded by the starvation stimulon, was more frequent in LTBI than in active TB (57). Collectively, these results strongly indicate that antigen recognition by T cells tracks antigen profiles expressed by tubercle bacilli in association with growth phase. It is worth mentioning, however, that one study found no difference in the response to ␣-crystallin between active TB and LTBI (64). One possible explanation is that strong differences in absolute bacillary load between latent infection and multibacillary forms of active TB overcome the effect of growth-phase-associated antigen expression. Cellular responses and disease progression. Very few studies have investigated the evolution of cellular immune responses over time. One such study showed that antigen- and mitogen-induced IFN-␥/IL-10 ratios were higher in household

CLIN. MICROBIOL. REV.

contacts of TB cases who did not develop disease than in those who did (68). In a more common study design, baseline levels of immune markers have been analyzed in relation to onset of disease at a later date. In these studies, higher baseline levels of IFN-␥ measured by IGRAs correlated with increased risk of disease progression (31, 37, 63, 84, 98). Similar results were obtained when tuberculin skin test responses were investigated (5). Thus, there is a correlation between levels of antigenspecific responses and bacillary burden, as proposed largely on the basis of animal studies (5). However, it has also been reported that increased IFN-␥ and decreased IL-4 production by ␣␤ and ␥␦ CD8⫹ T cells negatively correlate with development of disease in health care workers (105a). These results seem applicable to the general population, since similar conclusions were drawn in comparisons between subjects with LTBI and active TB patients (120a, 165). The basis of the apparent contradiction between the two sets of conclusions reported above is unclear, though it may lie in the cytokineexpressing T-cell population examined. The ratio between the Th2 cytokine IL-4 and the IL-4 splice variant IL-4␦2, which exhibits an expression pattern similar to that of IFN-␥ (165), has been proposed as another indicator of TB reactivation risk (38a, 162). Collectively, the results of these studies indicate that the imbalance between proinflammatory and anti-inflammatory responses associated with disease progression is reflected in the peripheral blood. The levels of immune markers can also change with severity of active disease. In one study, severe tuberculosis was associated with reduced IFN-␥ production by antigen-stimulated PBMCs (139), while in another study the number of antigenspecific T cells was higher in cavitary disease than in noncavitary disease (118a). The observed divergence likely results from the use of different antigens and assay types in the two studies. Humoral Immune Responses and Infection State While studies of T-cell immunity in TB have most typically focused on small numbers of immunodominant antigens, as mentioned above, the antibody response has been explored in relation to many antigenic targets. The evaluation of many antibody specificities has provided the opportunity to extensively assess the relationship between bacillary (i.e., antigen) burden and antibody response. Various aspects of this relationship are reviewed in this section, with an emphasis on results obtained with the serological interrogation of the full proteome of M. tuberculosis. We also review longitudinal studies investigating antibody responses during progression of the infection. Since the antibody response varies substantially from one patient to another, we also discuss the potential causes of antibody heterogeneity in tuberculosis. Antibody responses and bacterial metabolic status. A recent interrogation of the M. tuberculosis proteome (⬃4,000 proteins) with ⬃500 sera from TB suspects from countries where TB is endemic showed that approximately 10% of the bacterial proteome generates human antibody responses (76). These results define the immunoproteome, which contains predominantly membrane-associated and secreted proteins. Within the immunoproteome, a much smaller pool of proteins (⬍1% of the proteome) were preferentially recognized by sera from

VOL. 24, 2011

active TB patients. These were predominantly secreted proteins. These conclusions agree with much of the earlier serological work utilizing culture filtrates and purified secreted proteins (75a, 138, 139a, 142, 163a). The immunoproteome data strongly suggest that membrane-associated proteins (which might derive from low numbers of live bacilli, dead bacilli, or macrophage-secreted exosomes) are occasionally targeted during latent infection or paucibacillary disease. In either condition, the extracellular proteins are underrepresented, either because dormant bacilli do not secrete (latent infection) or because the numbers of metabolically active (and secreting) mycobacteria are low (paucibacillary disease). As bacillary burden increases with disease, metabolically active bacilli secrete proteins, which become the favored targets. Thus, antibody responses are dynamic and reflect the bacterial metabolic state during infection. Antibody responses and bacterial burden. The strongest evidence that antibody responses reflect bacillary burden derives from the many studies showing that antibody responses tend to be much stronger in sputum smear-positive than in smearnegative pulmonary TB (13a, 138, 142, 167a). The positive correlation between antibody levels and bacillary load was first described with use of purified or semipurified proteins, such as the 38-kDa antigen and Ag85 (13a, 167a), and it has now been seen at the immunoproteome level, indicating this to be a general characteristic of the antibody response in TB. With a few antigens, such as the 19-kDa lipoprotein, antibody levels have been reported as being higher in sera from smear-negative TB patients than in those from smear-positive patients, perhaps in correlation with particular HLA phenotypes (13). Hypotheses linking antigen recognition by antibody and HLA have not been tested. The strong correlation between antibody levels and bacillary burden constitutes a double-edged sword with regard to the usefulness of antibody as a biomarker of active TB. On one hand, conditions associated with low bacillary burden, such as stable asymptomatic infection and Mycobacterium bovis BCG vaccination, are by and large seronegative. Thus, antibodies can differentiate active TB from asymptomatic infection states that require no medical intervention. Moreover, seroconversion could be used as an early indication of disease progression, since activation of TB is presumably accompanied by an increased bacillary burden (see below). On the other hand, antibody-based assays have performed poorly when used to diagnose sputum smear-negative TB (1, 112, 138, 142), suggesting that antibody responses are ill-suited as markers of paucibacillary forms of active TB. A correlation between bacillary burden and antibody responses is also suggested by the observation that some subjects with a history of past TB are seropositive (29a, 76). These subjects may harbor a larger (or metabolically more active) bacterial population than the general latently infected population, as suggested by the observed association of past TB with increased risk of TB reactivation (58a, 104). While suggesting the intriguing possibility that increased antibody levels may help identify past TB cases in the process of reactivating TB, the occasional seropositivity seen in the past TB group poses an additional challenge to serodiagnosis of active TB. Antibody responses and disease progression. The association between the antibody response, bacillary load, and bacte-

IMMUNODIAGNOSIS OF TUBERCULOSIS

799

rial metabolic state suggests that antibody responses track disease progression. This possibility was tested in macaques, which respond to experimental M. tuberculosis infection with approximately equal probabilities of asymptomatic containment (latent infection) and active disease (15, 86). Moreover, some asymptomatic monkeys undergo spontaneous reactivation. It was found that in asymptomatic animals, antibody levels remained at preinfection levels or returned to preinfection levels after a transient increase (S. Kunnath, J. L. Flynn, and M. L. Gennaro, unpublished results). In contrast, antibody responses to the M. tuberculosis proteome increased in animals exhibiting active disease. The rise of antibody levels occurred at later times in the spontaneously reactivating animals relative to those classified as having acute active disease. Moreover, the number of antigenic targets increased with antibody levels in active disease, indicating that the number of antigens reaching threshold concentration levels for immune activation increases with antigen load. The findings in macaques agree with results of studies conducted on HIV-infected cohorts, where the levels of some antibodies were seen to increase prior to the diagnosis of active TB (16a, 50a, 76a, 138a). The fact that antibodies to some antigens but not to others increase with progression to disease (50a) again points to threshold levels for antibody production varying among immunodominant antigens. Whether these differences are attributable to regulatory mechanisms remains to be investigated. Diversity of antibody responses. The serum antibody profiles obtained from active TB patients differ from each other in terms of antigenic targets and titers. This has seriously hampered the development of TB serological diagnosis (as reviewed in many meta-analyses [1, 112, 142, 143]), since the requirement for multiple antigens as diagnostic reagents to increase the number of true-positive test results has been typically accompanied by increased false-positive test results. Host factors are almost certainly at play in determining serological diversity, since macaques infected by the same route with the same number of tubercle bacilli from the same strain develop varied antibody profiles (Kunnath et al., unpublished results). Moreover, antibodies from different murine strains recognize different antigen sets (69). In humans, the levels of some antibodies have been reported to be associated with HLA type (13, 13b). Variation may also be introduced by infecting bacterial strains differing in antigen composition. Variation most likely occurs at the level of expression of immunodominant antigens among clinical isolates (48a, 113a), since little diversity exists in genes encoding antigenic targets (102) and most antigenic epitopes are hyperconserved (22). For antigens in the PE/PPE family, strain-to-strain variation of gene expression has been proposed to provide tubercle bacilli with a dynamic antigenic profile (161). In addition to “intrinsic” host- and pathogen-derived factors, relative antigen burden can be viewed as a main source of antibody variability. It is likely that variability among antibody profiles results from the bacterial load and bacterial metabolic state at the time of testing and the relative immunodominance of each protein. Thus, the relative frequency at which the antibody response “samples” each immunodominant antigen will vary from one patient to another. Moreover, depending on relative antibody avidity, the effect of antigen load on the frequency of sampling will be greater for some antigens than for others. Furthermore, due to the chronic

800

KUNNATH-VELAYUDHAN AND GENNARO

nature of tuberculosis, these events occur over considerable periods of time; thus, cross-sectional analyses may further accentuate variation. It should also be noted that, given that a small subset of secretory proteins are specifically targeted during active TB (76), antibody responses to tuberculosis may appear homogeneous (128) under particular patient selection and testing conditions.

NEW PRINCIPLES FOR BIOMARKER DISCOVERY The following set of conclusions emerges from the work described above. 1. Clinically, infection with M. tuberculosis does not have a simple, binary outcome, i.e., latent infection and active disease. Each condition covers a spectrum of “subconditions.” Latent infection may be a stable state, it may be associated with a high risk of progression to disease, or it may represent a preclinical stage of disease. Active disease may be minimal, i.e., asymptomatic or accompanied by low-grade symptoms, or it may exhibit various degrees of severity in terms of symptoms, bacterial burden, and tissue damage. Thus, M. tuberculosis infection presents with a spectrum of multiple, often poorly separated, clinical conditions. 2. Histopathologically, the tuberculous granuloma is a dynamic structure. Host-pathogen interactions in the granuloma over the course of infection lead to adaptive changes of tubercle bacilli, of the phenotypes of the host immune cells, and of the levels of the immune mediators they produce. Recirculation of the immune cells and release of soluble mediators establish a link between local and systemic immune compartments. 3. Immunologically, the levels of some immune markers vary during infection because their expression is directly linked to immune function (e.g., protective or suppressive) and its regulation, others vary because they reflect changes in bacterial antigen composition and bacterial burden during infection, and yet others vary for both reasons. These considerations lead to the view that the spectrum of the clinical manifestations is accompanied by a corresponding spectrum of tissue damage and immune responses. If so, particular stages of the M. tuberculosis infection are associated with specific cellular phenotypes, cytokine levels, and/or antibody profiles. However, the association between biomarkers and infection stages is not simple, because it results from the interaction/intersection of multiple covariates that are host and pathogen derived (as it can be inferred from Fig. 2). Moreover, genetic and epigenetic diversity exists among hosts and among infecting strains, which further complicates the picture. Elements of diversity can be found even within the same host, since not all granulomas evolve at the same time and bacteria can be found at different growth phases within the same granuloma. Thus, for immunodiagnosis to be an effective tool for TB control, at least two conditions must be met. One is that immune markers associated with each particular clinical condition are found, and the other is that (at least some of) these markers are robust enough to withstand the heterogeneity associated with host- and pathogen-derived sources of variation.

CLIN. MICROBIOL. REV.

A Paradigm Shift The notion that M. tuberculosis infection is associated with a spectrum of ill-separated clinical conditions has become increasingly clear (see references 6 and 162 for two recent reports). Given that the host immune response is at the heart of TB pathogenesis, the presence of a concurrent immunological spectrum is almost evident. However, biomarker research has often shied away from the complexity of the immune response to M. tuberculosis infection, implementing instead a “reductionist” approach aiming at finding a marker (or marker set) that would diagnose “infection, yes/no” or “active TB, yes/no.” As a result, reactivity to an “active TB, yes” marker in the absence of active disease has been taken as a “false-positive” result rather than as the consequence of a simplistic case classification. Underestimating the consequences of the immunological spectrum and trying to force boundaries between states where boundaries barely exist seem to constitute an example of “forcing a square peg into a round hole.” It is our view that TB immunodiagnostic research is moving toward an inevitable paradigm shift, in which the presence (and even the absence) of immune markers exhibiting various degrees of association with a particular infection state provides an immune signature or “code” that is much more predictive of a particular infection state than any of the components of the code separately. Moreover, it may be expected that a complex signature is more robust than single markers with respect to person-to-person variation. Changes in the code (or a different code) would reflect transitions from one infection state to another. Systems Immunology The realization that no single marker identifies a particular M. tuberculosis infection stage with adequate diagnostic accuracy has led to the search for multiple markers. Thus far, this principle has been applied to markers of the same kind. For example, since serological recognition of M. tuberculosis antigens varies among TB patients, serodiagnostic research has been oriented toward multiantigen tests to reach suitable diagnostic sensitivity. Moreover, IGRAs for the diagnosis of LTBI have included two or three antigens for better sensitivity, even though large proportions of infected individuals exhibit cellular responses to immunodominant antigens such as ESAT-6 and CFP-10. However, the failure of serology to provide accurate diagnostics for active TB and of the current IGRAs to reliably distinguish between stable and progressive LTBI raises the possibility that the immunological signature of each M. tuberculosis infection state rests on a combination of immune markers of different types. For example, detectable antibody levels are strongly associated with active TB; however, the need for multiple antibodies to boost sensitivity reduces diagnostic specificity due to accumulating positive results in the population without TB disease. In contrast, the cellular response detected by the current IGRAs can be considered “almost universal” in infected individuals. It is not infrequent, however, that responses to IGRAs wane in TB patients, due to the immunosuppression associated with active disease. It is conceivable, therefore, that the diagnostic association of antibody to active disease might be strengthened by the concurrent absence of a particular cellular response that is

VOL. 24, 2011

IMMUNODIAGNOSIS OF TUBERCULOSIS

reduced in active disease due to immunoregulatory mechanisms. Additional diagnostic insight should result from chemokines or cytokines produced by peripheral blood cells when infection is contained rather than when immune control has failed. Moreover, it is conceivable that relative ratios of IgG isotypes, which reflect the cytokine environment, may also skew the diagnostic decision toward the presence or absence of an active disease process. All of the above could be further refined by taking into account antigen specificity of the responses, given the possibility that production of immunodominant antigens changes with the growth phase of the infecting bacilli. At least two alternative approaches can be envisioned for finding the diagnostic immune signatures of M. tuberculosis infection states. The first would take advantage of the large body of data sketched in the sections above. In this approach, combinatorial biomarkers would be identified from the investigation of known immune responses to known bacterial antigens. An alternative approach would take advantage of highthroughput technologies, which can generate information on thousands of “conditions” at once. Such technologies are already mature for some immune markers (e.g., antibodies to proteins and peptides). For others, such as the simultaneous detection of antigen-specific T-cell responses (58b, 103a), they are still being developed. High-throughput methods should make it feasible to characterize the immunological spectrum of M. tuberculosis infection by assessing large numbers of mediators of the cellular and humoral response to many (or all) antigens of the tubercle bacillus. Either approach would have to be used in longitudinal studies to assess how transitions from one infection state to another are associated with qualitative and quantitative changes in immune markers. The paradigm shift and the systems approach proposed above should also apply to pediatric TB, which differs from TB in adults with regard to risk of disease progression, pathophysiology, and clinical presentation (27). However, children infected or diseased with M. tuberculosis might express different immunological biomarkers than their adult counterparts, because critical differences exist between the innate and acquired responses of young children and adults (82) and because pediatric TB is often paucibacillary (141). CONCLUSIONS Characterizing the immunological spectrum of M. tuberculosis infection for immunodiagnostic purposes requires retooling TB immunodiagnostic research to better accommodate a dynamic, rather than static, view of biomarker discovery. The challenges are many. Longitudinal studies require large sample sizes, lengthy data collection periods, and costly resources. High-throughput methods are usually expensive. Assessment of combinatorial markers requires sophisticated analytical methods and extensive computing resources. The downstream development of multianalyte diagnostic assays that interrogate diverse components of the immune responses constitutes an area of research in its own right, particularly if the need for point-of-care tests drives the field toward lab-on-chip methodologies. However, each challenge can be turned into a new opportunity for research on TB immunodiagnostics, as was discussed at an international conference in 2008 (50). If the

801

will exists, with the help of international funding, advocacy, and political collaboration, these challenges can be met. The success will be worth the effort because, besides LTBI diagnosis by IGRAs, our current knowledge warns us that unless we break the code, no substantial advance will be made in immunodiagnosis for TB. ACKNOWLEDGMENTS We thank Yuri Bushkin, Karl Drlica, and Richard Pine for critical reading of the manuscript and C&M Consulting, Denville, NJ, for help with Fig. 2. Research in the Gennaro laboratory has been supported by the National Institutes of Health, the Foundation for New and Innovative Diagnostics, the European Union, and the Futura Foundation. REFERENCES 1. Abebe, F., C. Holm-Hansen, H. G. Wiker, and G. Bjune. 2007. Progress in serodiagnosis of Mycobacterium tuberculosis infection. Scand. J. Immunol. 66:176–191. 2. American Thoracic Society. 2000. Diagnostic standards and classification of tuberculosis in adults and children. Am. J. Respir. Crit. Care Med. 161: 1376–1395. 3. American Thoracic Society. 2000. Targeted tuberculin testing and treatment of latent tuberculosis infection. MMWR Recommend. Rep. 49:1–51. 4. Andersen, P., D. Askgaard, L. Ljungqvist, J. Bennedsen, and I. Heron. 1991. Proteins released from Mycobacterium tuberculosis during growth. Infect. Immun. 59:1905–1910. 5. Andersen, P., T. M. Doherty, M. Pai, and K. Weldingh. 2007. The prognosis of latent tuberculosis: can disease be predicted? Trends Mol. Med. 13:175– 182. 6. Barry, C. E., III, et al. 2009. The spectrum of latent tuberculosis: rethinking the biology and intervention strategies. Nat. Rev. Microbiol. 7:845–855. 7. Barry, S., R. Breen, M. Lipman, M. Johnson, and G. Janossy. 2009. Impaired antigen-specific CD4(⫹) T lymphocyte responses in cavitary tuberculosis. Tuberculosis (Edinb.) 89:48–53. 8. Beatty, W. L., et al. 2000. Trafficking and release of mycobacterial lipids from infected macrophages. Traffic 1:235–247. 9. Belisle, J. T., et al. 1997. Role of the major antigen of Mycobacterium tuberculosis in cell wall biogenesis. Science 276:1420–1422. 10. Bhatnagar, S., K. Shinagawa, F. J. Castellino, and J. S. Schorey. 2007. Exosomes released from macrophages infected with intracellular pathogens stimulate a proinflammatory response in vitro and in vivo. Blood 110:3234– 3244. 11. Biselli, R., et al. 2010. Detection of interleukin-2 in addition to interferongamma discriminates active tuberculosis patients, latently infected individuals, and controls. Clin. Microbiol. Infect. 16:1282–1284. 12. Boom, W. H. 1999. Gammadelta T cells and Mycobacterium tuberculosis. Microbes Infect. 1:187–195. 13. Bothamley, G. H., G. M. Schreuder, R. R. de Vries, and J. Ivanyi. 1993. Association of antibody responses to the 19-kDa antigen of Mycobacterium tuberculosis and the HLA-DQ locus. J. Infect. Dis. 167:992–993. 13a.Bothamley, G. H., R. Rudd, F. Festenstein, and J. Ivanyi. 1992. Clinical value of the measurement of Mycobacterium tuberculosis specific antibody in pulmonary tuberculosis. Thorax 47:270–275. 13b.Bothamley, G. H., et al. 1989. Association of tuberculosis and M. tuberculosis-specific antibody levels with HLA. J. Infect. Dis. 159:549–555. 14. Caccamo, N., et al. 2010. Multifunctional CD4(⫹) T cells correlate with active Mycobacterium tuberculosis infection. Eur. J. Immunol. 40:2211– 2220. 15. Capuano, S. V., III, et al. 2003. Experimental Mycobacterium tuberculosis infection of cynomolgus macaques closely resembles the various manifestations of human M. tuberculosis infection. Infect. Immun. 71:5831–5844. 16. Casey, R., et al. 2010. Enumeration of functional T-cell subsets by fluorescence-immunospot defines signatures of pathogen burden in tuberculosis. PLoS One 5:e15619. 16a.Cavalcante, S., et al. 1997. Association between an early humoral response to Mycobacterium tuberculosis antigens and later development of tuberculosis in human immunodeficiency virus-infected individuals. Int. J. Tuberc. Lung Dis. 1:170–174. 17. Chee, C. B., T. M. Barkham, K. W. Khinmar, S. H. Gan, and Y. T. Wang. 2009. Quantitative T-cell interferon-gamma responses to Mycobacterium tuberculosis-specific antigens in active and latent tuberculosis. Eur. J. Clin. Microbiol. Infect. Dis. 28:667–670. 18. Chee, C. B., et al. 2008. Comparison of sensitivities of two commercial gamma interferon release assays for pulmonary tuberculosis. J. Clin. Microbiol. 46:1935–1940. 19. Chegou, N. N., G. F. Black, M. Kidd, P. D. van Helden, and G. Walzl. 2009.

802

KUNNATH-VELAYUDHAN AND GENNARO

Host markers in QuantiFERON supernatants differentiate active TB from latent TB infection: preliminary report. BMC Pulm. Med. 9:21. 20. Chen, J., et al. 2009. Novel recombinant RD2- and RD11-encoded Mycobacterium tuberculosis antigens are potential candidates for diagnosis of tuberculosis infections in BCG-vaccinated individuals. Microbes Infect. 11: 876–885. 21. Clemens, D. L., B. Y. Lee, and M. A. Horwitz. 2002. The Mycobacterium tuberculosis phagosome in human macrophages is isolated from the host cell cytoplasm. Infect. Immun. 70:5800–5807. 22. Comas, I., et al. 2010. Human T cell epitopes of Mycobacterium tuberculosis are evolutionarily hyperconserved. Nat. Genet. 42:498–503. 23. Comstock, G. W., C. Baum, and D. E. Snider, Jr. 1979. Isoniazid prophylaxis among Alaskan Eskimos: a final report of the bethel isoniazid studies. Am. Rev. Respir. Dis. 119:827–830. 24. Connolly, L. E., P. H. Edelstein, and L. Ramakrishnan. 2007. Why is long-term therapy required to cure tuberculosis? PLoS Med. 4:e120. 25. Cooper, A. M. 2009. Cell-mediated immune responses in tuberculosis. Annu. Rev. Immunol. 27:393–422. 26. Covert, B. A., J. S. Spencer, I. M. Orme, and J. T. Belisle. 2001. The application of proteomics in defining the T cell antigens of Mycobacterium tuberculosis. Proteomics 1:574–586. 27. Cruz, A. T., and J. R. Starke. 2010. Pediatric tuberculosis. Pediatr. Rev. 31:13–26. 28. Cunningham, A. F., and C. L. Spreadbury. 1998. Mycobacterial stationary phase induced by low oxygen tension: cell wall thickening and localization of the 16-kilodalton alpha-crystallin homolog. J. Bacteriol. 180:801–808. 29. Daniel, T. M., M. J. Oxtoby, E. Pinto, and E. Moreno. 1981. The immune spectrum in patients with pulmonary tuberculosis. Am. Rev. Respir. Dis. 123:556–559. 29a.Davidow, A., et al. 2005. Antibody profiles characteristic of Mycobacterium tuberculosis infection state. Infect. Immun. 73:6846–6851. 30. Deb, C., et al. 2009. A novel in vitro multiple-stress dormancy model for Mycobacterium tuberculosis generates a lipid-loaded, drug-tolerant, dormant pathogen. PLoS One 4:e6077. 31. del Corral, H., et al. 2009. IFNgamma response to Mycobacterium tuberculosis, risk of infection and disease in household contacts of tuberculosis patients in Colombia. PLoS One 4:e8257. 32. Delves, P., S. Martin, D. Burton, and I. Roitt (ed.). 2006. Roitt’s essential immunology. Blackwell Publishing, Malden, MA. 33. Demissie, A., et al. 2006. Recognition of stage-specific mycobacterial antigens differentiates between acute and latent infections with Mycobacterium tuberculosis. Clin. Vaccine Immunol. 13:179–186. 34. Derrick, S. C., I. M. Yabe, A. Yang, and S. L. Morris. 2011. Vaccineinduced anti-tuberculosis protective immunity in mice correlates with the magnitude and quality of multifunctional CD4 T cells. Vaccine 29:2902– 2909. 35. Dheda, K., et al. 2008. Gene expression of IL17 and IL23 in the lungs of patients with active tuberculosis. Thorax 63:566–568. 36. Diel, R., et al. 2011. Interferon-gamma release assays for the diagnosis of latent Mycobacterium tuberculosis infection: a systematic review and metaanalysis. Eur. Respir. J. 37:88–99. 37. Diel, R., R. Loddenkemper, S. Niemann, K. Meywald-Walter, and A. Nienhaus. 2011. Negative and positive predictive value of a whole-blood interferon-␥ release assay for developing active tuberculosis: an update. Am. J. Respir. Crit. Care Med. 183:88–95. 38. Diel, R., R. Loddenkemper, and A. Nienhaus. 2010. Evidence-based comparison of commercial interferon-gamma release assays for detecting active TB: a metaanalysis. Chest 137:952–968. 38a.Doherty, M., R. S. Wallis, A. Zumia, and the WHO-Tropical Disease Research/European Commission Joint Expert Consultation Group. 2009. Biomarkers for tuberculosis disease status and diagnosis. Curr. Opin. Pulm. Med. 15:181–187. 39. Dominguez, J., et al. 2009. T-cell responses to the Mycobacterium tuberculosis-specific antigens in active tuberculosis patients at the beginning, during, and after antituberculosis treatment. Diagn. Microbiol. Infect. Dis. 63:43–51. 40. Dominguez, J., et al. 2008. Comparison of two commercially available gamma interferon blood tests for immunodiagnosis of tuberculosis. Clin. Vaccine Immunol. 15:168–171. 41. Dorhoi, A., S. T. Reece, and S. H. Kaufmann. 2011. For better or for worse: the immune response against Mycobacterium tuberculosis balances pathology and protection. Immunol. Rev. 240:235–251. 42. Dye, C., C. J. Watt, D. M. Bleed, S. M. Hosseini, and M. C. Raviglione. 2005. Evolution of tuberculosis control and prospects for reducing tuberculosis incidence, prevalence, and deaths globally. JAMA 293:2767–2775. 43. Ehlers, S. 2009. Lazy, dynamic or minimally recrudescent? On the elusive nature and location of the mycobacterium responsible for latent tuberculosis. Infection 37:87–95. 44. Eichbaum, Q., and E. J. Rubin. 2002. Tuberculosis. Advances in laboratory diagnosis and drug susceptibility testing. Am. J. Clin. Pathol. 118(Suppl.): S3–S17.

CLIN. MICROBIOL. REV. 45. Fenhalls, G., et al. 2002. Localisation of mycobacterial DNA and mRNA in human tuberculous granulomas. J. Microbiol. Methods 51:197–208. 46. Fenhalls, G., et al. 2002. Distribution of IFN-gamma, IL-4 and TNF-alpha protein and CD8 T cells producing IL-12p40 mRNA in human lung tuberculous granulomas. Immunology 105:325–335. 47. Fenhalls, G., et al. 2002. In situ detection of Mycobacterium tuberculosis transcripts in human lung granulomas reveals differential gene expression in necrotic lesions. Infect. Immun. 70:6330–6338. 48. Flynn, J. L., and J. Chan. 2001. Immunology of tuberculosis. Annu. Rev. Immunol. 19:93–129. 48a.Gao, Q., et al. 2005. Gene expression diversity among Mycobacterium tuberculosis clinical isolates. Microbiology 151:5–14. 49. Garton, N. J., et al. 2008. Cytological and transcript analyses reveal fat and lazy persister-like bacilli in tuberculous sputum. PLoS Med. 5:e75. 50. Gennaro, M. L., and T. M. Doherty (ed.). 2010. Immunodiagnosis of Tuberculosis: New questions, New Tools conference, 2008. Proceedings. BMC Proc. 4(Suppl. 3). http://www.tb-conference.com. 50a.Gennaro, M. L., et al. 2007. Antibody markers of incident tuberculosis among HIV-infected adults in the USA: a historical prospective study. Int. J. Tuberc. Lung Dis. 11:624–631. 51. Giri, P. K., and J. S. Schorey. 2008. Exosomes derived from M. Bovis BCG infected macrophages activate antigen-specific CD4⫹ and CD8⫹ T cells in vitro and in vivo. PLoS One 3:e2461. 52. Goletti, D., et al. 2010. Response to Rv2628 latency antigen associates with cured tuberculosis and remote infection. Eur. Respir. J. 36:135–142. 53. Goletti, D., et al. 2006. Accuracy of an immune diagnostic assay based on RD1 selected epitopes for active tuberculosis in a clinical setting: a pilot study. Clin. Microbiol. Infect. 12:544–550. 54. Goletti, D., et al. 2010. IFN-gamma, but not IP-10, MCP-2 or IL-2 response to RD1 selected peptides associates to active tuberculosis. J. Infect. 61:133– 143. 55. Goletti, D., et al. 2010. Is IP-10 an accurate marker for detecting M. tuberculosis-specific response in HIV-infected persons? PLoS One 5:e12577. 56. Goletti, D., et al. 2005. Selected RD1 peptides for active tuberculosis diagnosis: comparison of a gamma interferon whole-blood enzyme-linked immunosorbent assay and an enzyme-linked immunospot assay. Clin. Diagn. Lab. Immunol. 12:1311–1316. 57. Govender, L., et al. 2010. Higher human CD4 T cell response to novel Mycobacterium tuberculosis latency associated antigens Rv2660 and Rv2659 in latent infection compared with tuberculosis disease. Vaccine 29:51–57. 58. Green, A. M., et al. 2010. CD4(⫹) regulatory T cells in a cynomolgus macaque model of Mycobacterium tuberculosis infection. J. Infect. Dis. 202:533–541. 58a.Grzybowski, S. H. Fishaut, J. Rowe, and A. Brown. 1971. Tuberculosis among patients with various radiologic abnormalities, followed by the chest clinic service. Am. Rev. Respir. Dis. 104:605–608. 58b.Hadrup, S. R., et al. 2009. Parallel detection of antigen-specific T-cell responses by multidimensional encoding of MHC multimers. Nat. Methods 6:520–526. 59. Harari, A., et al. 2011. Dominant TNF-alpha(⫹) Mycobacterium tuberculosis-specific CD4(⫹) T cell responses discriminate between latent infection and active disease. Nat. Med. 17:372–376. 60. Hasan, Z., et al. 2009. Differential live Mycobacterium tuberculosis-, M. bovis BCG-, recombinant ESAT6-, and culture filtrate protein 10-induced immunity in tuberculosis. Clin. Vaccine Immunol. 16:991–998. 61. Hegazy, A. N., et al. 2010. Interferons direct Th2 cell reprogramming to generate a stable GATA-3(⫹)T-bet(⫹) cell subset with combined Th2 and Th1 cell functions. Immunity 32:116–128. 62. Hernandez-Pando, R., et al. 2000. Persistence of DNA from Mycobacterium tuberculosis in superficially normal lung tissue during latent infection. Lancet 356:2133–2138. 63. Higuchi, K., N. Harada, K. Fukazawa, and T. Mori. 2008. Relationship between whole-blood interferon-gamma responses and the risk of active tuberculosis. Tuberculosis (Edinb.) 88:244–248. 64. Hinks, T. S., et al. 2009. Frequencies of region of difference 1 antigenspecific but not purified protein derivative-specific gamma interferon-secreting T cells correlate with the presence of tuberculosis disease but do not distinguish recent from remote latent infections. Infect. Immun. 77:5486– 5495. 65. Hrabec, E., M. Strek, M. Zieba, S. Kwiatkowska, and Z. Hrabec. 2002. Circulation level of matrix metalloproteinase-9 is correlated with disease severity in tuberculosis patients. Int. J. Tuberc. Lung Dis. 6:713–719. 66. Humphrey, D. M., and M. H. Weiner. 1987. Mycobacterial antigen detection by immunohistochemistry in pulmonary tuberculosis. Hum. Pathol. 18:701–708. 67. Hussain, R., F. Shahid, S. Zafar, M. Dojki, and H. M. Dockrell. 2004. Immune profiling of leprosy and tuberculosis patients to 15-mer peptides of Mycobacterium leprae and M. tuberculosis GroES in a BCG vaccinated area: implications for development of vaccine and diagnostic reagents. Immunology 111:462–471.

VOL. 24, 2011 68. Hussain, R., N. Talat, F. Shahid, and G. Dawood. 2007. Longitudinal tracking of cytokines after acute exposure to tuberculosis: association of distinct cytokine patterns with protection and disease development. Clin. Vaccine Immunol. 14:1578–1586. 69. Huygen, K., et al. 1993. Influence of genes from the major histocompatibility complex on the antibody repertoire against culture filtrate antigens in mice infected with live Mycobacterium bovis BCG. Infect. Immun. 61:2687– 2693. 70. Jafari, C., et al. 2006. Rapid diagnosis of smear-negative tuberculosis by bronchoalveolar lavage enzyme-linked immunospot. Am. J. Respir. Crit. Care Med. 174:1048–1054. 71. Jafari, C., et al. 2008. Local immunodiagnosis of pulmonary tuberculosis by enzyme-linked immunospot. Eur. Respir. J. 31:261–265. 72. Jafari, C., and C. Lange. 2008. Suttons’s law: local immunodiagnosis of tuberculosis. Infection 36:510–514. 73. Janssens, J. P., et al. 2007. Quantitative scoring of an interferon-gamma assay for differentiating active from latent tuberculosis. Eur. Respir. J. 30:722–728. 74. Kanunfre, K. A., O. H. Leite, M. I. Lopes, M. Litvoc, and A. W. Ferreira. 2008. Enhancement of diagnostic efficiency by a gamma interferon release assay for pulmonary tuberculosis. Clin. Vaccine Immunol. 15:1028–1030. 75. Kaplan, G., et al. 2003. Mycobacterium tuberculosis growth at the cavity surface: a microenvironment with failed immunity. Infect. Immun. 71:7099– 7108. 75a.Kumar, G., et al. 2008. Diagnostic potential of Ag85C in comparison to various secretory antigens for childhood tuberculosis. Scand. J. Immunol. 68:177–183. 76. Kunnath-Velayudhan, S., et al. 2010. Dynamic antibody responses to the Mycobacterium tuberculosis proteome. Proc. Natl. Acad. Sci. U. S. A. 107:14703–14708. 76a.Laal, S., et al. 1997. Surrogate marker of preclinical tuberculosis in human immunodeficiency virus infection: antibodies to an 88-kDa secreted antigen of Mycobacterium tuberculosis. J. Infect. Dis. 176:133–143. 77. Lalvani, A., and M. Pareek. 2010. A 100 year update on diagnosis of tuberculosis infection. Br. Med. Bull. 93:69–84. 78. Lange, C., M. Pai, F. Drobniewski, and G. B. Migliori. 2009. Interferongamma release assays for the diagnosis of active tuberculosis: sensible or silly? Eur. Respir. J. 33:1250–1253. 79. Latorre, I., et al. 2009. Quantitative evaluation of T-cell response after specific antigen stimulation in active and latent tuberculosis infection in adults and children. Diagn. Microbiol. Infect. Dis. 65:236–246. 80. Lee, J. Y., et al. 2006. Comparison of two commercial interferon-gamma assays for diagnosing Mycobacterium tuberculosis infection. Eur. Respir. J. 28:24–30. 81. Lenzini, L., P. Rottoli, and L. Rottoli. 1977. The spectrum of human tuberculosis. Clin. Exp. Immunol. 27:230–237. 82. Lewinsohn, D. A., M. L. Gennaro, L. Scholvinck, and D. M. Lewinsohn. 2004. Tuberculosis immunology in children: diagnostic and therapeutic challenges and opportunities. Int. J. Tuberc. Lung Dis. 8:658–674. 83. Leyten, E. M., et al. 2006. Human T-cell responses to 25 novel antigens encoded by genes of the dormancy regulon of Mycobacterium tuberculosis. Microbes Infect. 8:2052–2060. 84. Lienhardt, C., et al. 2010. Evaluation of the prognostic value of IFN-gamma release assay and tuberculin skin test in household contacts of infectious tuberculosis cases in Senegal. PLoS One 5:e10508. 85. Lighter, J., M. Rigaud, M. Huie, C. H. Peng, and H. Pollack. 2009. Chemokine IP-10: an adjunct marker for latent tuberculosis infection in children. Int. J. Tuberc. Lung Dis. 13:731–736. 86. Lin, P. L., et al. 2009. Quantitative comparison of active and latent tuberculosis in the cynomolgus macaque model. Infect. Immun. 77:4631–4642. 87. Ling, D. I., L. L. Flores, L. W. Riley, and M. Pai. 2008. Commercial nucleic-acid amplification tests for diagnosis of pulmonary tuberculosis in respiratory specimens: meta-analysis and meta-regression. PLoS One 3:e1536. 88. Ling, D. I., A. A. Zwerling, and M. Pai. 2008. GenoType MTBDR assays for the diagnosis of multidrug-resistant tuberculosis: a meta-analysis. Eur. Respir. J. 32:1165–1174. 89. Macedo, G. C., A. Bozzi, H. R. Weinreich, A. Bafica, H. C. Teixeira, and S. C. Oliveira. 2011. Human T cell and antibody-mediated responses to the Mycobacterium tuberculosis recombinant 85A, 85B, and ESAT-6 antigens. Clin. Dev. Immunol. 2011:351573. 90. Maglione, P. J., and J. Chan. 2009. How B cells shape the immune response against Mycobacterium tuberculosis. Eur. J. Immunol. 39:676–686. 91. Mase, S. R., et al. 2007. Yield of serial sputum specimen examinations in the diagnosis of pulmonary tuberculosis: a systematic review. Int. J. Tuberc. Lung Dis. 11:485–495. 92. Masungi, C., et al. 2002. Differential T and B cell responses against Mycobacterium tuberculosis heparin-binding hemagglutinin adhesin in infected healthy individuals and patients with tuberculosis. J. Infect. Dis. 185:513–520. 93. McDonough, K. A., Y. Kress, and B. R. Bloom. 1993. Pathogenesis of

IMMUNODIAGNOSIS OF TUBERCULOSIS

803

tuberculosis: interaction of Mycobacterium tuberculosis with macrophages. Infect. Immun. 61:2763–2773. 94. Mendez, A., R. Hernandez-Pando, S. Contreras, D. Aguilar, and G. A. Rook. 2011. CCL2, CCL18 and sIL-4R in renal, meningeal and pulmonary TB; a 2 year study of patients and contacts. Tuberculosis (Edinb.) 91:140– 145. 95. Menzies, D. 2008. Using tests for latent tuberculous infection to diagnose active tuberculosis: can we eat our cake and have it too? Ann. Intern. Med. 148:398–399. 96. Menzies, D., G. Gardiner, M. Farhat, C. Greenaway, and M. Pai. 2008. Thinking in three dimensions: a web-based algorithm to aid the interpretation of tuberculin skin test results. Int. J. Tuberc. Lung Dis. 12:498–505. 97. Menzies, D., M. Pai, and G. Comstock. 2007. Meta-analysis: new tests for the diagnosis of latent tuberculosis infection: areas of uncertainty and recommendations for research. Ann. Intern. Med. 146:340–354. 98. Metcalfe, J. Z., et al. 2010. Evaluation of quantitative IFN-gamma response for risk stratification of active tuberculosis suspects. Am. J. Respir. Crit. Care Med. 181:87–93. 99. Millington, K. A., et al. 2007. Dynamic relationship between IFN-gamma and IL-2 profile of Mycobacterium tuberculosis-specific T cells and antigen load. J. Immunol. 178:5217–5226. 100. Montoya, D., and R. L. Modlin. 2010. Learning from leprosy: insight into the human innate immune response. Adv. Immunol. 105:1–24. 101. Mukamolova, G. V., O. Turapov, J. Malkin, G. Woltmann, and M. R. Barer. 2010. Resuscitation-promoting factors reveal an occult population of tubercle bacilli in sputum. Am. J. Respir. Crit. Care Med. 181:174–180. 102. Musser, J. M., A. Amin, and S. Ramaswamy. 2000. Negligible genetic diversity of mycobacterium tuberculosis host immune system protein targets: evidence of limited selective pressure. Genetics 155:7–16. 103. Nemeth, J., et al. 2009. Recruitment of Mycobacterium tuberculosis specific CD4⫹ T cells to the site of infection for diagnosis of active tuberculosis. J. Intern. Med. 265:163–168. 103a.Newell, E. W., L. O. Klein, W. Yu, and M. M. Davis. 2009. Simultaneous detection of many T-cell specificities using combinatorial tetramer staining. Nat. Methods 6:497–499. 104. Nolan, C. M., and A. M. Elarth. 1988. Tuberculosis in a cohort of Southeast Asian refugees. A five-year surveillance study. Am. Rev. Respir. Dis. 137: 805–809. 105. North, R. J., and Y. J. Jung. 2004. Immunity to tuberculosis. Annu. Rev. Immunol. 22:599–623. 105a.Ordway, D. J., et al. 2004. Increased interleukin-4 production by CD8 and gammadelta T cells in health-care workers is associated with the subsequent development of active tuberculosis. J. Infect. Dis. 190:756–766. 106. Ottenhoff, T. H. M., D. A. Lewinsohn, and D. M. Lewinsohn. 2008. Human CD4 and CD8 T cell responses to Mycobacterium tuberculosis: antigen specificity, function, implications and applications, p. 119–155. In S. H. E. Kaufmann and W. J. Britton (ed.), Handbook of tuberculosis: immunology and cell biology. Wiley-VCH Verlag GmbH & co KGaA, Weinheim, Germany. 107. Pai, M. 2010. Spectrum of latent tuberculosis—existing tests cannot resolve the underlying phenotypes. Nat. Rev. Microbiol. 8:242. (Author reply, 8:242.) 108. Pai, M., K. Dheda, J. Cunningham, F. Scano, and R. O’Brien. 2007. T-cell assays for the diagnosis of latent tuberculosis infection: moving the research agenda forward. Lancet Infect. Dis. 7:428–438. 109. Pai, M., L. W. Riley, and J. M. Colford, Jr. 2004. Interferon-gamma assays in the immunodiagnosis of tuberculosis: a systematic review. Lancet Infect. Dis. 4:761–776. 110. Pai, M., A. Zwerling, and D. Menzies. 2008. Systematic review: T-cell-based assays for the diagnosis of latent tuberculosis infection: an update. Ann. Intern. Med. 149:177–184. 111. Pantaleo, G., and A. Harari. 2006. Functional signatures in antiviral T-cell immunity for monitoring virus-associated diseases. Nat. Rev. Immunol. 6:417–423. 112. Parkash, O., B. P. Singh, and M. Pai. 2009. Regions of differences encoded antigens as targets for immunodiagnosis of tuberculosis in humans. Scand. J. Immunol. 70:345–357. 113. Pathan, A. A., et al. 2001. Direct ex vivo analysis of antigen-specific IFNgamma-secreting CD4 T cells in Mycobacterium tuberculosis-infected individuals: associations with clinical disease state and effect of treatment. J. Immunol. 167:5217–5225. 113a.Pheiffer, C., J. C. Betts, H. R. Flynn, P. T. Lukey, and P. van Helden. 2005. Protein expression by a Beijing strain differs from that of another clinical isolate and Mycobacterium tuberculosis H37Rv. Microbiology 151:1139– 1150. 114. Pollock, J. M., and P. Andersen. 1997. The potential of the ESAT-6 antigen secreted by virulent mycobacteria for specific diagnosis of tuberculosis. J. Infect. Dis. 175:1251–1254. 115. Price, N. M., R. H. Gilman, J. Uddin, S. Recavarren, and J. S. Friedland. 2003. Unopposed matrix metalloproteinase-9 expression in human tuberculous granuloma and the role of TNF-alpha-dependent monocyte networks. J. Immunol. 171:5579–5586.

804

KUNNATH-VELAYUDHAN AND GENNARO

116. Rachman, H., et al. 2006. Unique transcriptome signature of Mycobacterium tuberculosis in pulmonary tuberculosis. Infect. Immun. 74:1233–1242. 117. Ravn, P., et al. 1999. Human T cell responses to the ESAT-6 antigen from Mycobacterium tuberculosis. J. Infect. Dis. 179:637–645. 118. Ravn, P., et al. 2005. Prospective evaluation of a whole-blood test using Mycobacterium tuberculosis-specific antigens ESAT-6 and CFP-10 for diagnosis of active tuberculosis. Clin. Diagn. Lab Immunol. 12:491–496. 118a.Ribeiro, S., et al. 2009. T-SPOT.TB responses during treatment of pulmonary tuberculosis. BMC Infect. Dis. 9:23. 119. Ridley, D. S. 1974. Histological classification and the immunological spectrum of leprosy. Bull. World Health Organ. 51:451–465. 120. Roberts, D. M., R. P. Liao, G. Wisedchaisri, W. G. Hol, and D. R. Sherman. 2004. Two sensor kinases contribute to the hypoxic response of Mycobacterium tuberculosis. J. Biol. Chem. 279:23082–23087. 120a.Roberts, T., N. Beyers, A. Aguirre, and G. Walzl. 2007. Immunosuppression during active tuberculosis is characterized by decreased interferon-gamma production and CD25 expression with elevated forkhead box P3, transforming growth factor-beta, and interleukin-4 mRNA levels. J. Infect. Dis. 195: 870–888. 121. Rogerson, B. J., et al. 2006. Expression levels of Mycobacterium tuberculosis antigen-encoding genes versus production levels of antigen-specific T cells during stationary level lung infection in mice. Immunology 118:195– 201. 122. Rook, G. A. 2007. Th2 cytokines in susceptibility to tuberculosis. Curr. Mol. Med. 7:327–337. 123. Rueda, C. M., N. D. Marin, L. F. Garcia, and M. Rojas. 2010. Characterization of CD4 and CD8 T cells producing IFN-gamma in human latent and active tuberculosis. Tuberculosis (Edinb.) 90:346–353. 124. Ruhwald, M., M. Bjerregaard-Andersen, P. Rabna, J. Eugen-Olsen, and P. Ravn. 2009. IP-10, MCP-1, MCP-2, MCP-3, and IL-1RA hold promise as biomarkers for infection with M. tuberculosis in a whole blood based T-cell assay. BMC Res. Notes 2:19. 125. Ruhwald, M., et al. 2007. CXCL10/IP-10 release is induced by incubation of whole blood from tuberculosis patients with ESAT-6, CFP10 and TB7.7. Microbes Infect. 9:806–812. 126. Russell, D. G. 2007. Who puts the tubercle in tuberculosis? Nat. Rev. Microbiol. 5:39–47. 127. Russell, D. G., P. J. Cardona, M. J. Kim, S. Allain, and F. Altare. 2009. Foamy macrophages and the progression of the human tuberculosis granuloma. Nat. Immunol. 10:943–948. 128. Samanich, K., J. T. Belisle, and S. Laal. 2001. Homogeneity of antibody responses in tuberculosis patients. Infect. Immun. 69:4600–4609. 129. Sargentini, V., et al. 2009. Cytometric detection of antigen-specific IFNgamma/IL-2 secreting cells in the diagnosis of tuberculosis. BMC Infect. Dis. 9:99. 130. Schaible, U. E., et al. 2003. Apoptosis facilitates antigen presentation to T lymphocytes through MHC-I and CD1 in tuberculosis. Nat. Med. 9:1039– 1046. 131. Schuck, S. D., et al. 2009. Identification of T-cell antigens specific for latent Mycobacterium tuberculosis infection. PLoS One 4:e5590. 132. Schwander, S., and K. Dheda. 2011. Human lung immunity against Mycobacterium tuberculosis: insights into pathogenesis and protection. Am. J. Respir. Crit. Care Med. 183:696–707. 133. Seder, R. A., P. A. Darrah, and M. Roederer. 2008. T-cell quality in memory and protection: implications for vaccine design. Nat. Rev. Immunol. 8:247– 258. 134. Sester, M., et al. 2011. Interferon-gamma release assays for the diagnosis of active tuberculosis: a systematic review and meta-analysis. Eur. Respir. J. 37:100–111. 135. Shi, L., Y. J. Jung, S. Tyagi, M. L. Gennaro, and R. J. North. 2003. Expression of Th1-mediated immunity in mouse lungs induces a Mycobacterium tuberculosis transcription pattern characteristic of nonreplicating persistence. Proc. Natl. Acad. Sci. U. S. A. 100:241–246. 136. Shi, L., R. North, and M. L. Gennaro. 2004. Effect of growth state on transcription levels of genes encoding major secreted antigens of Mycobacterium tuberculosis in the mouse lung. Infect. Immun. 72:2420–2424. 137. Shi, L., et al. 2005. Changes in energy metabolism of Mycobacterium tuberculosis in mouse lung and under in vitro conditions affecting aerobic respiration. Proc. Natl. Acad. Sci. U. S. A. 102:15629–15634. 138. Silva, V. M., G. Kanaujia, M. L. Gennaro, and D. Menzies. 2003. Factors associated with humoral response to ESAT-6, 38 kDa and 14 kDa in patients with a spectrum of tuberculosis. Int. J. Tuberc. Lung Dis. 7:478– 484. 138a.Simonney, N., et al. 2007. B-cell immune responses in HIV positive and HIV negative patients with tuberculosis evaluated with an ELISA using a glycolipid antigen. Tuberculosis (Edinb.) 87:109–122. 139. Sodhi, A., J. Gong, C. Silva, D. Qian, and P. F. Barnes. 1997. Clinical correlates of interferon gamma production in patients with tuberculosis. Clin. Infect. Dis. 25:617–620. 139a.Sonnenberg, M. G., and J. T. Belisle. 1997. Definition of Mycobacterium tuberculosis culture filtrate proteins by two-dimensional polyacrylamide gel

CLIN. MICROBIOL. REV. electrophoresis, N-terminal amino acid sequencing, and electrospray mass spectrometry. Infect. Immun. 65:4515–4524. 140. Sousa, A. O., et al. 1998. IgG subclass distribution of antibody responses to protein and polysaccharide mycobacterial antigens in leprosy and tuberculosis patients. Clin. Exp. Immunol. 111:48–55. 141. Starke, J. R. 2007. New concepts in childhood tuberculosis. Curr. Opin. Pediatr. 19:306–313. 142. Steingart, K. R., et al. 2009. Performance of purified antigens for serodiagnosis of pulmonary tuberculosis: a meta-analysis. Clin. Vaccine Immunol. 16:260–276. 143. Steingart, K. R., et al. 2007. Commercial serological antibody detection tests for the diagnosis of pulmonary tuberculosis: a systematic review. PLoS Med. 4:e202. 144. Steingart, K. R., et al. 2006. Fluorescence versus conventional sputum smear microscopy for tuberculosis: a systematic review. Lancet Infect. Dis. 6:570–581. 145. Steingart, K. R., et al. 2006. Sputum processing methods to improve the sensitivity of smear microscopy for tuberculosis: a systematic review. Lancet Infect. Dis. 6:664–674. 146. Strassburg, A., C. Jafari, M. Ernst, W. Lotz, and C. Lange. 2008. Rapid diagnosis of pulmonary TB by BAL enzyme-linked immunospot assay in an immunocompromised host. Eur. Respir. J. 31:1132–1135. 147. Sutherland, J. S., I. M. Adetifa, P. C. Hill, R. A. Adegbola, and M. O. Ota. 2009. Pattern and diversity of cytokine production differentiates between Mycobacterium tuberculosis infection and disease. Eur. J. Immunol. 39: 723–729. 148. Sutherland, J. S., B. C. de Jong, D. J. Jeffries, I. M. Adetifa, and M. O. Ota. 2010. Production of TNF-alpha, IL-12(p40) and IL-17 can discriminate between active TB disease and latent infection in a West African cohort. PLoS One 5:e12365. 149. Syed Ahamed Kabeer, B., B. Raman, A. Thomas, V. Perumal, and A. Raja. 2010. Role of QuantiFERON-TB gold, interferon gamma inducible protein-10 and tuberculin skin test in active tuberculosis diagnosis. PLoS One 5:e9051. 150. Teitelbaum, R., et al. 1999. Mycobacterial infection of macrophages results in membrane-permeable phagosomes. Proc. Natl. Acad. Sci. U. S. A. 96: 15190–15195. 151. Teitelbaum, R., et al. 1998. A mAb recognizing a surface antigen of Mycobacterium tuberculosis enhances host survival. Proc. Natl. Acad. Sci. U. S. A. 95:15688–15693. 152. Timm, J., et al. 2003. Differential expression of iron-, carbon-, and oxygenresponsive mycobacterial genes in the lungs of chronically infected mice and tuberculosis patients. Proc. Natl. Acad. Sci. U. S. A. 100:14321–14326. 153. Torrado, E., and A. M. Cooper. 2010. IL-17 and Th17 cells in tuberculosis. Cytokine Growth Factor Rev. 21:455–462. 154. Tsao, T. C., et al. 2002. Shifts of T4/T8 T lymphocytes from BAL fluid and peripheral blood by clinical grade in patients with pulmonary tuberculosis. Chest 122:1285–1291. 155. Tully, G., et al. 2005. Highly focused T cell responses in latent human pulmonary Mycobacterium tuberculosis infection. J. Immunol. 174:2174– 2184. 156. Ulrichs, T., and S. H. Kaufmann. 2006. New insights into the function of granulomas in human tuberculosis. J. Pathol. 208:261–269. 157. Ulrichs, T., et al. 2005. Differential organization of the local immune response in patients with active cavitary tuberculosis or with nonprogressive tuberculoma. J. Infect. Dis. 192:89–97. 158. Ulrichs, T., et al. 2004. Human tuberculous granulomas induce peripheral lymphoid follicle-like structures to orchestrate local host defence in the lung. J. Pathol. 204:217–228. 159. van der Wel, N., et al. 2007. M. tuberculosis and M. leprae translocate from the phagolysosome to the cytosol in myeloid cells. Cell 129:1287–1298. 160. Voskuil, M. I., et al. 2003. Inhibition of respiration by nitric oxide induces a Mycobacterium tuberculosis dormancy program. J. Exp. Med. 198:705– 713. 161. Voskuil, M. I., K. C. Visconti, and G. K. Schoolnik. 2004. Mycobacterium tuberculosis gene expression during adaptation to stationary phase and low-oxygen dormancy. Tuberculosis (Edinb.) 84:218–227. 162. Wallis, R. S., et al. 2010. Biomarkers and diagnostics for tuberculosis: progress, needs, and translation into practice. Lancet 375:1920–1937. 163. Walzl, G., K. Ronacher, W. Hanekom, T. J. Scriba, and A. Zumla. 2011. Immunological biomarkers of tuberculosis. Nat. Rev. Immunol. 11:343– 354. 163a.Wang, B. L., et al. 2005. Antibody response to four secretory proteins from Mycobacterium tuberculosis and their complex antigen in TB patients. Int. J. Tuberc. Lung Dis. 9:1327–1334. 164. Wang, J. Y., et al. 2007. Diagnosis of tuberculosis by an enzyme-linked immunospot assay for interferon-gamma. Emerg. Infect. Dis. 13:553–558. 165. Wassie, L., et al. 2008. Ex vivo cytokine mRNA levels correlate with changing clinical status of Ethiopian TB patients and their contacts over time. PLoS One 3:e1522. 166. Wayne, L. G., and C. D. Sohaskey. 2001. Nonreplicating persistence of Mycobacterium tuberculosis. Annu. Rev. Microbiol. 55:139–163.

VOL. 24, 2011

IMMUNODIAGNOSIS OF TUBERCULOSIS

805

167. Whittaker, E., A. Gordon, and B. Kampmann. 2008. Is IP-10 a better biomarker for active and latent tuberculosis in children than IFNgamma? PLoS One 3:e3901. 167a.Wiker, H. G., and M. Harboe. 1992. The antigen 85 complex: a major secretion product of Mycobacterium tuberculosis. Microbiol. Rev. 56:648–661. 168. Wilkinson, K. A., et al. 2005. Ex vivo characterization of early secretory antigenic target 6-specific T cells at sites of active disease in pleural tuberculosis. Clin. Infect. Dis. 40:184–187. 169. Wolf, A. J., et al. 2008. Initiation of the adaptive immune response to

Mycobacterium tuberculosis depends on antigen production in the local lymph node, not the lungs. J. Exp. Med. 205:105–115. 170. Yoshiyama, T., N. Harada, K. Higuchi, Y. Sekiya, and K. Uchimura. 2010. Use of the QuantiFERON-TB Gold test for screening tuberculosis contacts and predicting active disease. Int. J. Tuberc. Lung Dis. 14:819– 827. 171. Young, J. M., I. M. Adetifa, M. O. Ota, and J. S. Sutherland. 2010. Expanded polyfunctional T cell response to mycobacterial antigens in TB disease and contraction post-treatment. PLoS One 5:e11237.

Shajo Kunnath-Velayudhan, M.B.B.S., M.M.S.T., obtained his medical degree from the Trichur Medical College, Kerala, India, and his master’s degree in medical science and technology from the Indian Institute of Technology, Kharagpur, India. As a postdoctoral fellow in the Gennaro laboratory, he studied proteome-scale antibody responses to Mycobacterium tuberculosis infection.

Maria Laura Gennaro, M.D., is a Professor of Medicine at the Public Health Research Institute, New Jersey Medical School, Newark, NJ. Over the years, she has investigated bacterial pathogens, including enterotoxic Escherichia coli, Salmonella spp., Vibrio cholerae, Staphylococcus aureus, and Mycobacterium tuberculosis. The main areas of Dr. Gennaro’s tuberculosis research have been the antibody response to M. tuberculosis infection and the remodeling of the M. tuberculosis transcriptome during infection. She is currently leading a systems biology approach to the study of M. tuberculosis physiology during infection and of the interaction between the tubercle bacillus and the host macrophage.