Microbiology Case Study: a 53 Year Old Man with a Black Spot on His Shoulder

Case History

A 53 year old man presents to urgent care with a primary complaint of an area of erythema and tenderness around a small black spot on his left shoulder, shortly after returning from Ecuador. He does not report any fevers, chills, or drainage from the affected area. The patient reported that he occasionally felt the area moving. An occlusive Vaseline dressing was applied to the central black spot, and the organism shown below emerged from the wound.

Laboratory Identification

The parasite shown above is a human botfly larva, Dermatobia hominis. The clinical history is strongly suspicious for a botfly infection, and the patient himself suggested the diagnosis.

Dermatobia hominis is identified in large part by its relatively unique presentation combined with identification of the larvae in tissue. Laboratory identification of genus and species involves comparing morphological structures including the anterior and posterior spiracles, mouthparts and cephalopharyngeal skeleton, and cuticular spines. Travel history can also be helpful for genus or species-level identification.

Discussion

The lifecycle of human botflies begins when the female botfly lays her eggs on a mosquito. Once a mosquito feeds on a host, the botfly larva drop onto the host and burrow into the skin. They may remain in that location for up to 10 weeks before dropping off the host into soil to pupate and continue the life cycle.

The human botfly is found in North America, ranging from Mexico to Paraguay and northeast Argentina. Cases in the US are primarily in travelers returning from the botfly’s native range. Measuring the incidence of infection in travelers can be difficult due to the nature of the disease. Experienced travelers may be able to remove the larva at home. In other cases the botfly larva may leave the host before the patient seeks medical care.

Testing for the presence of these larva is easy as they require oxygen coming in through a hole in the skin. Cover the lesion with a thick layer of sterile Vaseline gauze and wait approximately 5-15 minutes for the organism to emerge. Surgery is usually not required as the larva is most often removed intact. Antibiotics should be given following removal of the parasite to prevent secondary infections.

-Britt Boles, MD is a 1st year Anatomic and Clinical Pathology Resident at the University of Vermont Medical Center.

-Christi Wojewoda, MD, is the Director of Clinical Microbiology at the University of Vermont Medical Center and an Associate Professor at the University of Vermont.

Microbiology Case Study: A 27 Year Old with Disseminated Joint Pain

Case History

A 27 year old male presented to the Emergency Department (ED) with complaints of right knee pain and swelling for one week. Two weeks prior, he tripped while walking to work and began to feel pain in his right calf. Upon physical examination, swelling was noted in his ankles, knee, shoulders, and fingers. The knee and shoulder were tender to palpation. In the ED, he was afebrile and vitals were normal. He denied any sort of injury, chills, or rash and no history of tobacco, alcohol, or illicit substance abuse. CT scan of the lower extremity showed no acute fracture but moderate to large knee joint effusion was observed. He and his fiancé (male partner) has been in a monogamous relationship for almost a decade, however the patient did have a history of gonorrhea nine years ago but was treated. Knee arthrocentesis was performed. The fluid was yellow and cloudy and contained 27,000 WBCs. The Gram stain of the synovial fluid showed many intracellular gram negative diplococci and the joint fluid culture grew out Neisseria gonorrhoeae. PCR of the rectal swab also detected N. gonorrhoeae.

Discussion

N. gonorrhoeae is the causative agent of gonorrhea, a sexually transmitted disease. In the United States, it is the second most commonly reported communicable disease.1 While infections can be asymptomatic, in men, gonorrhea commonly causes acute urethritis with dysuria, urethral discharge, and rarely, epididymitis.2,3,4 In women, gonorrhea can cause cervicitis and lead to pelvic inflammatory disease (PID), infertility, ectopic pregnancy, and chronic pelvic pain.5,6 Those with gonococcal endocervicitis can be co-infected with Chlamydia trachomatis and/or Trichomonas vaginalis, other causative agents of sexually transmitted diseases. N. gonorrhoeae can cause extragenital infections in the pharynx and rectum, which are most commonly seen among men who have sex with men (MSM). Disseminated gonococcal infection is rare (0.5-3% of infected individuals) and can be characterized by low grade fever, hemorrhagic skin lesions, tenosynovitis, polyarthralgia and septic arthritis. Complications of disseminated infections may include permanent joint damage, endocarditis, and meningitis. Gonococcal conjunctivitis mainly affects newborns from untreated mothers.7

Gonorrhea can be diagnosed clinically by a history and physical examination and also, by microbiological methods. Home collection kits are available to increase convenience. On a Gram stain, N. gonorrhoeae, a gram negative coccus, frequently appears within or closely associated polymorphonuclear leukocytes (PMNs) typically as diplococci pairs. Direct smears can be prepared from urethral, endocervical sites, and normally sterile or minimally contaminated sites such as joint fluid. Swab specimens (e.g. urogenital, pharyngeal, vaginal or rectal) should be collected with a Dacron or Rayon swab as calcium alginate and cotton swabs may be toxic or inhibitory for the bacteria.8 Specimens must be transported to the microbiology immediately. 9 Blood and joint fluid are also acceptable specimen types for culture for detection of disseminated gonococcal infection.

Enriched selective media for culture of N. gonorrhoeae includes MTM medium, ML medium, GC-Lect and the New York City medium. Plates should be incubated in a CO2 incubator (between 3-7%) at 35C to 37C for optimal growth.9 Gram negative diplococci recovered from urogenital sites that grow on the selective media and are oxidase-positive can be presumptively identified as N. gonorrhoeae. Another quick biochemical test that can be done is superoxol; N. gonorrhoeae produce immediate bubbling whereas N. meningitidis and N. lactamica produce weak, delayed bubbling. Confirmation using other testing methods such as carbohydrate utilization tests (e.g. N. gonorrhoeae produces acid from glucose only), immunological methods, enzymatic procedures, or DNA probe are also available.10

Compared to standard culture methods, Nucleic Acid Amplification Tests (NAAT) offer more rapid results and increased sensitivity. Additionally, NAATs may also include additional targets such as C. trachomatis, a frequent co-pathogen, as part of the assay. NAATs should be used according manufacturer’s protocols and on validated specimen types. For example, the Cepheid Xpert CT/NG test (as used by our patient here) can be used to test asymptomatic and symptomatic individuals and the acceptable specimen types are urine, pharyngeal, and rectal swabs, patient-collected vaginal swabs, and clinician-collected endocervical swabs.11 Given the legal implications of a N. gonorrhoeae diagnosis in a child, the CDC recommends that NAATs can be used to test for N. gonorrhoeae from vaginal and urine specimens from females and urine for males.12 For extragenital specimens, only validated FDA-cleared NAATs assays using pediatric specimens should be used.

The CDC recommends that uncomplicated gonorrhea be treated with ceftriaxone and azithromycin. However, between 2000-2010s, elevated MICs to both ceftriaxone and cefixime were seen and emerging azithromycin resistance is still a concern. The CLSI M100 currently recommends agar dilution or disk diffusion for antimicrobial susceptibility testing for N. gonorrhoeae. Susceptible and resistant interpretative breakpoints are available for penicillin, most cephems, tetracycline, ciprofloxacin, and spectinomycin. Of note, for azithromycin, only the susceptible category has a breakpoint.13

Image 1. Gram stain of synovial fluid showing many intracellular gram negative diplococci.
Image 2. Chlamydia trachomatis and Neisseria gonorrhoeae PCR. Orange and Brown= targets for N. gonorrhoeae; light and dark green=control genes.

References

  1. CDC. Sexually Transmitted Disease Surveillance, 2020. Atlanta, GA: Department of Health and Human Services; April 2022.
  2. John J, Donald WH. Asymptomatic urethral gonorrhoea in men. Br J Vener Dis 1978; 54:322.
  3. Handsfield HH, Lipman TO, Harnisch JP, et al. Asymptomatic gonorrhea in men. Diagnosis, natural course, prevalence and significance. N Engl J Med 1974; 290:117.
  4. Sherrard J, Barlow D. Gonorrhoea in men: clinical and diagnostic aspects. Genitourin Med 1996; 72:422.
  5. McCormack WM, Johnson K, Stumacher RJ, Donner A, Rychwalski R. Clinical spectrum of gonococcal infection in women. Lancet, 1(8023), 1182–1185 (1977).
  6. Curran J, Rendtorff R, Chandler R, Wiser W, Robinson H. Female gonorrhea: its relation to abnormal uterine bleeding, urinary tract symptoms, and cervicitis. Obstet Gynecol, 45(2), 195–198 (1975).
  7. O’Brien JP, Goldenberg DL, Rice PA. Disseminated gonococcal infection: a prospective analysis of 49 patients and a review of pathophysiology and immune mechanisms. Medicine (Baltimore) 1983; 62:395.
  8. Laurer BA, Masters HB. Toxic effect of calcium alginate swabs on Neiserria gonorrhoeae. J Clin Microbiol 1988: 26:54-56
  9. Leber, A. 3.9 Genital Cultures. Clinical Microbiology Procedures Handbook, 4th Edition. ASM Press, Washington, DC. 2016. p.3.9.3.1-3.9.3.15. 
  10. Knapp JS. Historical perspectives and identification of Neisseria and related species. Clin Microbiol Rev 1988;1:415-431.
  11. Cepheid GeneXpert. Xpert CT/NG (English). 2019. 301-0234 Rev.K
  12. CDC. Gonococcal Infections Among Infants and Children. Sexually Transmitted Infection Treatment Guidelines, Atlanta, GA: Department of Health and Human Services; 2021.
  13. CLSI. Performance Standards for Antimicrobial Susceptibility Test. CLSI supplement M100. Wayne, PA: Clinical and Laboratory Standards Institute; 2022, Edition 32

-Maikel Benitez Barzaga, MD is a Pathology Resident (PGY-1) at The George Washington University Hospital. His academic interest include hematology, microbiology, molecular and surgical pathology.

-Rebecca Yee, PhD, D(ABMM), M(ASCP)CM is the Chief of Microbiology, Director of Clinical Microbiology and Molecular Microbiology Laboratory at the George Washington University Hospital. Her interests include bacteriology, antimicrobial resistance, and development of infectious disease diagnostics.

Microbiology Case Study: A 32 Year Old with Lower Extremity Swelling

Case History

A 32 year old male with alcoholic cirrhosis presented to the emergency department with progressive lower extremity swelling. On presentation he was found to have jaundice due to hemolytic anemia secondary to spur cell anemia. Admission hemoglobin was 4.3 mg/dL (4.0-11.0 mg/dL) and bilirubin, both total and direct, were 6.3 mg/dL (0.2-1.3 mg/dL) and 2.9 mg/dL (0.0-0.5 mg/dL), respectively. He also had acute kidney injury (AKI) thought to be secondary to hepatorenal syndrome leading to the development of anasarca. A urinalysis was performed as part of the evaluation for his AKI that showed 100 WBC/HPF, > 187 RBC/HPF, and moderate bacteria which triggered a urine culture.

Laboratory Identification

Urine received in the microbiology laboratory was plated on Blood and MacConkey/CNA agars and grew non-hemolytic, lactose-fermenting gram negative rods (Image 1). Indole testing was negative. Given this biochemical pattern, a member of the Enterobacterales was suspected as typically seen in urine cultures. However, MALDI-TOF MS provided the surprising identification of Salmonella enterica subsp. arizonae. Xylose Lysine Deoxycholate (XLD) agar was set up to confirm the unusual identification (Image 2). Hydrogen sulfide production is typical of Salmonellae, and lactose fermentation, a trait unique to some isolates of S. enterica subsp. arizonae, was confirmed. The organism was submitted to the Texas Department of Health laboratory where the isolate was definitively identified as Salmonella enterica subsp. arizonae (IIIa 14:z4,z23) by whole genome sequencing.

Image 1. Patient isolate of S. enterica subsp. arizonae exhibiting lactose fermentation on MacConkey agar after 18 hours of incubation at 35°C (A). Lactose-fermentation is a unique hallmark of S. enterica subsp. arizonae compared to other Salmonellae (B).
Image 2. Patient isolate of S. enterica subsp. arizonae exhibiting hydrogen sulfide production and lactose fermentation on XLD agar after 18 hours at 35°C (A). Note the abundant yellow color of the medium (black arrowhead) compared to S. enterica subsp. Enterica serovar Enteritidis which does not ferment lactose, but also produces hydrogen sulfide (B, white arrowhead).

Discussion

This is a rare case of an extraintestinal infection caused by Salmonella enterica subsp. arizonae. Salmonellaeare motile, gram negative, facultatively anaerobic bacilli that are members of the Enterobacterales. The genus is composed of two species, S. enterica and S. bongori. Salmonella enterica is further subdivided into six subspecies: enterica (group I), salamae (group II), arizonae (group IIIa), diarizonae (group IIIb), houtenae (group IV), and indica (group VI). Salmonella bongori used to be classified as group V but was separated as a unique species based on genomic analysis.1 S. bongori almost exclusively causes zoonotic infections, while S. enterica subsp. enterica is the most frequent cause of human clinical disease. Salmonella taxonomy is complicated further by the division of members of S. enterica subsp. enterica into >2500 unique serovars based on immunoreactivity to lipopolysaccharide (O) and two flagellar (H) surface antigens. These are then further separated into “typhoidal” and “non-typhoidal” serovars based upon the characteristics of infection (Image 3).

Image 3. Hierarchical structure of Salmonella taxonomy. S. enterica subsp. arizonae is boxed in red to highlight is taxonomic position away from other pathogenic Salmonellae. Adapted from reference number 6.

Until recently, determinative testing was almost uniformly performed by serological confirmation of agglutination with O and H antigen-specific antisera. This has been a mainstay of epidemiological analysis of foodborne Salmonella outbreaks. Only recently has whole genome sequencing been adapted as a higher throughput and more discriminatory alternative to classical serotyping schemes. Salmonella nomenclature often uses a genus-species-subspecies format followed by serovar (e.g. Salmonella enterica subsp. enterica serovar Typhi), or it can be reported as genus-serovar for short (e.g. Salmonella Typhi). Formal identification will include information concerning the two flagellar antigens and lipopolysaccharide antigens, in addition to the formalized subspecies using the formula: genus-species-subspecies [space] O antigens [colon] Phase 1 H antigen [comma] Phase 2 H antigen. In this case, the formal identification from the state laboratory for this isolate was Salmonella enterica subsp. arizonae IIIa 14:z4,z23.

About 99% of human infections are due to Salmonella enterica subspecies enterica (group I)including the serotypes Enteritidis, Typhimurium, Typhi, Paratyphi.2 Infections due to Salmonella enterica subspecies arizonae are rare; serovar IIIa 41:z4,z23 is associated with 10-20 infections per year.3 Infection typically begins as gastroenteritis from food poising or from animal sources, particularly reptiles or poultry. Disease is typically seen in the young and immunocompromised and can progress to invasive disease including sepsis, meningitis, and osteomyelitis.4 It is unclear why there are lower rates of Salmonella enterica subspecies arizonae infections in humans as compared to Salmonella enterica subspecies enterica, but there is evidence to suggest Salmonella enterica subspecies arizonae and diarizonae have altered intestinal colonization in murine models leading to failure of Salmonella to persist in the mammalian intestinal tract.5

This patient had alcoholic cirrhosis and uncomplicated cystitis secondary to Salmonella extraintestinal infection at the time of presentation. It is unclear if this patient had gastroenteritis prior to developing cystitis and the limited medical history did not reveal exposure to reptiles or poultry. In this case, the patient completed seven days of ceftriaxone without complication or recurrence of infection.

References

  1. Agbaje M, Begum RH, Oyekunle MA, Ojo OE, Adenubi OT. Evolution of Salmonella nomenclature: a critical note. Folia Microbiol (Praha) 2011; 56(6): 497-503.
  2. Brenner FW, Villar RG, Angulo FJ, Tauxe R, Swaminathan B. Salmonella nomenclature. J Clin Microbiol 2000; 38(7): 2465-7.
  3. Shariat NW, Timme RE, Walters AT. Phylogeny of Salmonella enterica subspecies arizonae by whole-genome sequencing reveals high incidence of polyphyly and low phase 1 H antigen variability. Microb Genom 2021; 7(2).
  4. Abbott SL, Ni FC, Janda JM. Increase in extraintestinal infections caused by Salmonella enterica subspecies II-IV. Emerg Infect Dis 2012; 18(4): 637-9.
  5. Katribe E, Bogomolnaya LM, Wingert H, Andrews-Polymenis H. Subspecies IIIa and IIIb Salmonellae are defective for colonization of murine models of salmonellosis compared to Salmonella enterica subsp. I serovar typhimurium. J Bacteriol 2009; 191(8): 2843-50.
  6. Achtman M, Wain J, Weill FX, Nair S, Zhou Z, et al. (2012) Multilocus Sequence Typing as a Replacement for Serotyping in Salmonella enterica. PLOS Pathogens 8(6): e1002776. https://doi.org/10.1371/journal.ppat.1002776

Denver Niles, MD is the Medical Microbiology fellow at UT Southwestern Medical Center. Prior to his Medical Microbiology fellowship, he completed pediatric infectious disease training at Baylor College of Medicine/Texas Children’s Hospital.

Muluye Mesfin, SM(ASCP)CM is the microbiology laboratory supervisor at UT Southwestern Medical Center where he has worked for 12 years.  Prior to this, Mo completed a bachelor of science degree in medical technology at the University of Maryland.

-Clare McCormick-Baw, MD, PhD is an Assistant Professor of Clinical Microbiology at UT Southwestern in Dallas, Texas. She has a passion for teaching about laboratory medicine in general and the best uses of the microbiology lab in particular.

-Andrew Clark, PhD, D(ABMM) is an Assistant Professor at UT Southwestern Medical Center in the Department of Pathology, and Associate Director of the Clements University Hospital microbiology laboratory. He completed a CPEP-accredited postdoctoral fellowship in Medical and Public Health Microbiology at National Institutes of Health, and is interested in antimicrobial susceptibility and anaerobe pathophysiology.

Microbiology Case Study: An Adult Woman with a Pelvic Abscess

Case History

An adult woman presented to the emergency department five days after undergoing gynecological surgery. The patient presented with fever and severe right lower quadrant abdominal pain. Computed tomography (CT) scan with contrast showed a ring enhanced loculated fluid collection within the cervix, which was concerning for an abscess. The patient was admitted to the hospital and empirically started on piperacillin-tazobactam, but continued to have fevers despite the antibiotics. Blood and urine samples were sent to the microbiology lab for bacterial culture but no organisms were isolated from either source. Two days later, the patient underwent a diagnostic laparoscopy, abdominal wash-out, and drainage of the abscess. The abscess fluid was sent for aerobic and anaerobic bacterial culture. Gram stain of the specimen showed 3+ white blood cells with no organism seen. The anaerobic culture grew 4+ pinpoint white colonies on blood agar after 5 days of incubation. Further identification of these colonies by MALDI-TOF MS revealed Mycoplasma hominis.

Image 1. Blood agar with 4+ pinpoint translucent colonies.

Discussion

Mycoplasma hominis is often a commensal of the urogenital tract, but it can be associated with urogenital infections including pelvic inflammatory disease (PID), pregnancy-related infections, and urethritis in males. There are multiple risk factors for Mycoplasma hominis genital infection including young adult age, multiple sexual partners, and pregnancy. Immunocompromised patients have a higher risk for Mycoplasma hominis extragenital infections as nearly 50% of reported extragenital infections isolated from immunocompromised patients.2 Mycoplasma hominis can cause extragenital infections including septic arthritis,4 septicemia, osteitis, retroperitoneal abscesses3, mediastinitis,1 and pneumonia.

Laboratory diagnosis of Mycoplasma hominis is challenging due to the fastidious nature of the organism and its lack of the cell wall makes it undetectable by gram staining. The more specific tests including molecular tests for Mycoplasma hominis are not routinely ordered unless there is a strong clinical suspicion, which makes diagnosis more challenging. Mycoplasma hominis can grow on 5% sheep blood and chocolate agars; however, such growth is very slow and may take from 2 to 7 days of incubation.1 The usual growth of Mycoplasma hominis reveals tiny-sized pinpoint colonies that may be overlooked (Image 1). Once growth is observed, MALDI-TOF MS can be used for identification.6

There are multiple types of selective media for the isolation of Mycoplasma hominis including SP4 agar supplemented with arginine, Hayflick agar, A7, and A8 agars.9 Both A7 and A8 agars contain arginine to enrich Mycoplasma growth but differ in the antibiotic content used to inhibit the growth of other commensals. Agar plates should be put for incubation under 5 to 10% CO2 or under anaerobic conditions at 35°C for at least 5 days.9 On these selective agars Mycoplasma hominis has a characteristic fried egg appearance and can be seen by the aid of a stereomicroscope. However, use of specific agar is not widespread.

Molecular testing of Mycoplasma hominis using nucleic acid amplification (NAAT) assays such as polymerase chain reaction (PCR) is a more sensitive and faster method of detecting Mycoplasma hominis compared with culture. However, PCR is neither widely available nor standardized. PCR assays for Mycoplasma hominis generally use 16S rRNA as a gene target, but other targets, including gap, fstY, and yidC, have been developed.7 Clinical picture should be taken into account when evaluating the significance of a positive PCR test as Mycoplasma hominis can be a commensal organism and PCR does not distinguish between live and dead organisms.

Mycoplasma spp. lack a peptidoglycan cell wall. This makes Mycoplasma spp. intrinsically resistant to β-lactams and to all antibiotics, which target the cell wall, including glycopeptide antibiotics. Mycoplasma hominis is also resistant to rifampin, sulfonamides and trimethoprim. Tetracyclines, macrolides, and fluoroquinolones are often used. Antimicrobial susceptibility testing is rarely performed, with only a few specialized laboratories offering the testing. Clinical and laboratory standards institute guidelines (CLSI M43) is followed using microbroth dilution. Agar disc diffusion testing is not used for Mycoplasma hominis as there is no correlation between inhibitory zones and minimal inhibitory concentrations.8 Mycoplasma hominis can be evaluated for susceptibility to clindamycin, tetracycline, and levofloxacin.10

After isolation of Mycoplasma hominis was reported, doxycycline was added to the patient’s antibiotic regimen. The patient responded well with subsiding of the fever and stabilization of her vital signs.

References

  1. Xiang, L., & Lu, B. 2019. Infection due to Mycoplasma hominis after left hip replacement: case report and literature review. BMC infectious diseases, 19(1), 50. https://doi.org/10.1186/s12879-019-3686-z
  2. Meyer RD, Clough W. 1993. Extragenital Mycoplasma hominis infections in adults: emphasis on immunosuppression. Clin Infect Dis. Suppl 1:S243-9. doi: 10.1093/clinids/17.supplement_1.s243. PMID: 8399923.
  • Adams M, Bouzigard R, Al-Obaidi M, Zangeneh TT. 2020. Perinephric abscess in a renal transplant recipient due to Mycoplasma hominis: Case report and review of the literature. Transpl Infect Dis.(5):e13308. doi: 10.1111/tid.13308. Epub 2020 Jul 7. PMID: 32378787.
  • Luttrell LM, Kanj SS, Corey GR, Lins RE, Spinner RJ, Mallon WJ, Sexton DJ. 1994. Mycoplasma hominis septic arthritis: two case reports and review. Clin Infect Dis.19(6):1067-70. doi: 10.1093/clinids/19.6.1067. PMID: 7888535.
  • Wylam ME, Kennedy CC, Hernandez NM, Peters SG, Maleszewski JJ, Cassivi SD, Scott JP. 2013. Fatal hyperammonemia caused by Mycoplasma hominis. Lancet 382:1956.
  • Pereyre S, Tardy F, Renaudin H, Cauvin E, Del Pra Netto Machado L, Tricot A, Benoit F, Treilles M, Bebear C. 2013. Identification and subtyping of clinically relevant human and ruminant mycoplasmas by use of matrix-assisted laser desorption ionization–time of flight mass spectrometry. J Clin Microbiol 51:3314–3323.
  • Ferandon C, Peuchant O, Janis C, Benard A, Renaudin H, Pereyre S, Bebear C. 2011. Development of a real-time PCR targeting the yidC gene for the detection of Mycoplasma hominis and comparison with quantitative culture. Clin Microbiol Infect 17:155–159.
  • Clinical and Laboratory Standards Institute. 2011. Methods for antimicrobial susceptibility testing for human mycoplasmas; approved guideline M43-A. Clinical and Laboratory Standards Institute, Wayne, PA.
  • Stabler S, Faure E, Duployez C, Wallet F, Dessein R, Le Guren R. 2021. Mycoplasma hominis extragenital abscess. J Clin Microbiol, 59(4). https://doi.org/10.1128/JCM.02343-20
  • https://sites.uab.edu/dml/tests/

Omar Abdelsadek, MD is a PGY-1 (AP/CP) Pathology Resident at University of Chicago (NorthShore) Pritzker School of Medicine.

-Paige M.K. Larkin, PhD, D(ABMM), M(ASCP)CM is the Director of Molecular Microbiology and Associate Director of Clinical Microbiology at NorthShore University HealthSystem in Evanston, IL. Her interests include mycology, mycobacteriology, point-of-care testing, and molecular diagnostics, especially next generation sequencing.

Microbiology Case Study: Worsening Liver Function and Bacteremia in a 35 Year Old Male

Case History

A 35 year old male with a history of alcohol use disorder in early remission, acute alcoholic hepatitis with multiple admissions for worsening liver function was admitted for acute kidney injury and worsening encephalopathy. Blood cultures were collected due to leukocytosis and the anaerobic bottle flagged positive for gram negative bacilli at 4.6 days. The organism, shown in Image 1, was sent to a reference laboratory and was identified as a Campylobacter species, unable to further identify. The patient will receive a liver transplant at another institution.

Image 1. Campylobacter species morphology in a blood smear.

Discussion

Campylobacter species are gram-negative, oxidase-positive, non-fermenting, microaerophilic, non-spore forming, motile rods typically with one or more helical turn.1,2 When two bacteria form short chains, these appear as “S” shaped and/or “gull-wing” shaped. These bacteria are generally 0.2 µm by 0.5-5.0 µm in size and can be as long as 8.0 µm.1 Campylobacter species are widely distributed in most warm-blooded animals (e.g., poultry, cattle, pigs, sheep, cats, and dogs) and they grow optimally at 37-42 °C. There are more than 20 Campylobacter species, not all of which cause illness but are potentially pathogenic. Campylobacter jejuni accounts for approximately 90% of human Campylobacter infections, while less common species such as Campylobacter coli, Campylobacter upsaliensis, Campylobacter fetus, and Campylobacter lari can also cause infection.3

Transmission of Campylobacter is believed to be foodborne via undercooked meat (particularly poultry), unpasteurized milk, or improperly treated water. Person-to-person transmission is rare, but may occur via the fecal-oral route. The infection load for Campylobacter species is relatively low, with fewer than 500 organisms causing infection.4 In human infection, these bacteria usually colonize the intestinal tract leading to diarrhea (often bloody), stomach cramps, fever, nausea, and vomiting.5 Clinical manifestation usually occurs 2 to 5 days after the individual is infected and lasts approximately a week. Diagnosis is established definitively by stool culture and sometimes by blood culture.2 In some cases, long-term effects of Campylobacter infection include an array of clinical syndromes including enteritis, bacteremia, arthritis, septic abortion, meningitis, irritable bowel disease, and Guillain-Barre syndrome [4]. Individuals with a greater risk for infection include those 65-years or older, pregnant women, and those with weakened immune systems.5

Campylobacteriosis is the most common form of acute infectious diarrhea in developed countries with a higher incidence than both Salmonella and Shigella.1 The Center for Disease Control and Prevention estimates that 1.5 million people in the United States are affected by Campylobacter infection each year—making it the most common bacterial cause of diarrheal illness in the United States.3 Unfortunately, the incidence of hepatitis associated with Campylobacter species infection is unknown, as few case-reports related to Campylobacter colitis6and Campylobacter jejuni 7,8,9,10 have been published. Although the liver is often involved in systemic infections resulting in various types of abnormal liver function tests, mild to severe hepatocellular dysfunction is an uncommon observation in those with Campylobacter infection.

Most individuals infected with any Campylobacter species recover with only fluid replenishment while the diarrhea lasts and no antibiotic treatment. However, those with or at risk for severe illness should be considered for antibiotic treatment. The antibiotics that are used to treat infection are azithromycin and fluoroquinolones (usually resistant). Antimicrobial susceptibility testing can help guide appropriate therapy.3

References

  1. Hardy Diagnostics. Campylobacter [Internet]. 2016. Available from: https://catalog.hardydiagnostics.com/cp_prod/Content/hugo/Campylobacter.htm#:~:text=In%20general%2C%20Campylobacter%20spp.%20appear%20as%20gray%2C%20flat%2C,glistening%2C%20with%20little%20spreading.%20Campylobacter%20spp.%20are%20non-hemolytic.
  2. Perez-Perez GI, Blaser MJ. Campylobacter and Helicobacter. In: Baron S, editor. Medical Microbiology. Galveston (TX): University of Texas Medical Branch at Galveston Copyright © 1996, The University of Texas Medical Branch at Galveston.; 1996.
  3. Centers for Disease Control and Prevention. Campylobacter (Campylobacteriosis) For Health Professionals [Internet]. 2019 [updated December 23, 2019]. Available from: https://www.cdc.gov/campylobacter/technical.html.
  4. Ehrenpreis ED. Campylobacter infection [Internet]. Epocrates2022 [updated January 22, 2022]. Available from: https://online.epocrates.com/v2/print/disease/1175?subSectionId=11#:~:text=Bacteria%20of%20the%20genus%20Campylobacter%20cause%20a%20variety,%5B%203%5D%20There%20are%20many%20species%20of%20Campylobacter.
  5. Centers for Disease Control and Prevention. Campylobacter (Campylobacteriosis) Symptoms [Internet]. 2019. Available from: https://www.cdc.gov/campylobacter/symptoms.html.
  6. Reddy KR, Farnum JB, Thomas E. Acute hepatitis associated with campylobacter colitis. J Clin Gastroenterol. 1983;5(3):259-62. Epub 1983/06/01. doi: 10.1097/00004836-198306000-00013. PubMed PMID: 6863882.
  7. Humphrey KS. Campylobacter infection and hepatocellular injury. Lancet. 1993;341(8836):49. Epub 1993/01/02. doi: 10.1016/0140-6736(93)92521-t. PubMed PMID: 8093289.
  8. Vermeij CG, van Dissel JT, Veenendaal RA, Lamers CB, van Hoek B. Campylobacter jejuni peritonitis in a patient with liver cirrhosis. Eur J Gastroenterol Hepatol. 1996;8(12):1219-21. Epub 1996/12/01. doi: 10.1097/00042737-199612000-00016. PubMed PMID: 8980944.
  9. Korman TM, Varley CC, Spelman DW. Acute hepatitis associated with Campylobacter jejuni bacteraemia. Eur J Clin Microbiol Infect Dis. 1997;16(9):678-81. Epub 1997/11/14. doi: 10.1007/bf01708559. PubMed PMID: 9352262.
  10. Yoon JG, Lee SN, Hyun HJ, Choi MJ, Jeon JH, Jung E, et al. Campylobacter jejuni Bacteremia in a Liver Cirrhosis Patient and Review of Literature: A Case Study. Infect Chemother. 2017;49(3):230-5. Epub 2017/06/14. doi: 10.3947/ic.2017.49.3.230. PubMed PMID: 28608661; PubMed Central PMCID: PMCPMC5620392

-Amelia M. Lamberty is a MS in Pathology student at the Larner College of Medicine at the University of Vermont.

-Christi Wojewoda, MD, is the Director of Clinical Microbiology at the University of Vermont Medical Center and an Associate Professor at the University of Vermont.

Microbiology Case Study: A 69 Year Old Man with Chronic Cutaneous Disease

Case Description

A 69 year old man with hepatitis B and chronic cutaneous Rosai-Dorfman disease presented to the dermatology clinic for regular follow-up. He was being treated with subcutaneous injection of methotrexate every other week and intralesional Kenalog (ILK) injections for individual lesions. The patient presented with a new complaint of a painful nodule on his left thumb where he was stuck with a splinter two months prior. He denied fever, chills, weight loss, or other systemic symptomology. Upon physical examination, an erythematous nodule on the lateral left thumb with central pallor and crusting consistent with a foreign body was observed (Figure 1). Surgical excision was recommended.

Figure 1. Photograph of a nodular lesion on the lateral aspect of the left thumb with surrounding erythema and central pallor, which was described as painful, and had been present for two months following traumatic splinter implantation.

Following excisional biopsy, histopathology revealed a relatively circumscribed lesion with suppurative granulomatous dermatitis and numerous pigmented hyphae observed on hematoxylin and eosin stained slides (H&E; [Figures 2-3]). A diagnosis of phaeohyphomycosis was made; the patient’s methotrexate was held and an infectious disease (ID) consult was placed. Precautionary blood cultures were drawn which remained negative following five days of incubation. The patient was started on a course of empiric oral doxycycline for two weeks which he completed. At presentation for ID follow up, the patient felt well and denied constitutional symptoms or recurrence of the thumb lesion. Physical exam revealed no associated sporotrichoid lesions (lymphocutaneous spread of infection) or palpable lymphadenopathy. ID recommended a 3-month course of oral itraconazole as secondary prophylaxis, which was completed without adverse effects or recurrence of symptoms.

Figure 2. Hematoxylin and Eosin (H&E) stained excisional biopsy demonstrating a relatively circumscribed lesion with suppurative granulomatous inflammation (100x magnification). Thick arrows highlight lavender staining epithelioid-histiocytes that comprise the bulk of the granuloma and arrowheads point to the admixed aggregates of dark pink and purple staining neutrophils, giving this granuloma its suppurative nature.
Figure 3. High-power magnification photomicrograph of the lesion with a close-up view of the suppurative nature of this granulomatous inflammation with small arrows highlighting the numerous cross-sections of hyphae that demonstrate melanized pigment observed in this case of phaeohyphomycosis (H&E stain; 400x magnification).

Case Discussion

Phaeohyphomycosis describes a constellation of clinical syndromes caused by infection with a broad group of “dematiaceous” or “melanized” molds and some pigmented yeasts.1 Many of these organisms are ubiquitous in the environment though some are more selective in their habitat, restricting the likelihood of infection to specific geography or select patient populations.2,3 Despite significant microbiological diversity, a unifying characteristic of dematiaceous molds is the production of the pigment melanin. Melanin is theorized to serve as a virulence factor, as loss of melanization often results in attenuation.3,4  In contrast to other diseases caused by dematiaceous molds with more defined etiologies and presentations (e.g., eumycetoma, chromoblastomycosis), manifestations of phaeohyphomycosis are highly variable and can include keratitis, cutaneous disease, pulmonary infection, central nervous system penetration and/or disseminated disease.

Laboratory diagnosis of phaeohyphomycosis is reliant on histopathological evaluation, as surgical debridement is often necessary for management. In this setting, darkly pigmented, septate hyphae invading tissue in a nonspecific background of inflammation may be observed.1 H&E staining is generally sufficient to confirm diagnosis; however, special stains that can highlight fungi, namely Grocott-Gomori methenamine silver (GMS) or periodic acid-Schiff (PAS) stains, can outline the presence of hyphal elements. Additionally, melanin production can be highlighted using Fontana-Masson staining. Careful evaluation and interpretation of fungal cultures, when collected, are important as results can be complex given the ubiquitous nature of many etiological agents, particularly from non-sterile anatomical sites. Additionally, there are no alternate methods routinely available to aid in diagnosis, outside of culture, to specifically identify etiologic agents of phaeohyphomycosis.3 Importantly, optimal antifungal therapy for these infections remains unclear due to a lack of randomized control trials and relative infrequency of presentation.

Superficial infections, such as the one described in this case, are generally considered to be consequences of local trauma, and exhibit minimal tissue invasion. However, in the setting of the immunocompromised host or immunosuppression, disseminated infection can occur.3 The prognosis of invasive phaeohyphomycosis is poor, exhibiting a mortality is as high as 10% for deep local infections and 50% for disseminated disease.1 This patient’s advanced age and chronic immunosuppression were cause for great concern. Fortunately, the biopsy demonstrated granuloma formation effectively localizing the infection to the subcutaneous tissue of the thumb. The patient has remained free of further disease to date, suggestive of a curative surgical resection.

References

  1. Arcobello, JT, Revankar, SG. Phaeohyphomycosis. Respiratory and Critical Care Medicine. 2020. DOI: 10.1055/s-0039-3400957
  2. Wong EH, Revankar SG. Dematiaceous Molds. Infectious Disease Clinics of North America. 2016. 10.1016/j.idc.2015.10.007
  3. Revankar, SG., Baddley, JW., Chen, S.C-A., Kauffman, CA., Slavin, M., Vazquez, JA, Seas, C., Morris, MI., Nguyen, MH et. al. A Mycoses Study Group International Prospective Study of Phaeohyphomycosis: and Analysis of 99 Proven/Probable Cases. Open Forum Infectious Diseases. 2017. DOI: 10.1093/ofid/ofx200
  4. Sharkey PK, Graybill JR, Rinaldi MG, Stevens DA, Tucker RM, Peterie JD, Hoeprich PD, Greer DL, Frenkel L, Counts GW, et al. Itraconazole treatment of phaeohyphomycosis. Journal of the American Academy of Dermatology. 1990. doi: 10.1016/0190-9622(90)70259-k

-Kevin Burningham is a 4th year medical student at UT Southwestern Medical School.

-Dominick Cavuoti is a Professor at UT Southwestern Medical Center who specializes in Cytopathology, Infectious Disease pathology and is a medical director of the Microbiology laboratory at Parkland Health and Hospital System.

-Clare McCormick-Baw, MD, PhD is an Assistant Professor of Clinical Microbiology at UT Southwestern in Dallas, Texas. She has a passion for teaching about laboratory medicine in general and the best uses of the microbiology lab in particular.

-Andrew Clark, PhD, D(ABMM) is an Assistant Professor at UT Southwestern Medical Center in the Department of Pathology, and Associate Director of the Clements University Hospital microbiology laboratory. He completed a CPEP-accredited postdoctoral fellowship in Medical and Public Health Microbiology at National Institutes of Health, and is interested in antimicrobial susceptibility and anaerobe pathophysiology.

Microbiology Case Study: An Elderly Women Presents with Altered Mental Status, Fever, and Gastrointestinal Symptoms

Case Description

An elderly woman presented to the emergency department with sudden onset diarrhea, fever and altered mental status for 1 day. On admission, she had fever with chills, tachycardia, tachypnea, and diminishing mental status. Her white blood cell (WBC) count was slightly elevated from baseline with neutrophilia and thrombocytopenia. Blood and CSF were sent for bacterial culture. Stool was sent for C. difficile PCR and occult blood, which came back negative.

Organisms were recovered from the aerobic blood culture bottle. Gram stain showed gram positive short rods. A multiplex PCR was run directly from the blood, which detected Listeria monocytogenes. The care team was notified immediately. The organism grew on blood agar and aerobic Columbia Naladixic Acid Agar (CNA agar) with characteristic beta hemolysis. The identification was re-confirmed by MALDI-TOF MS as Listeria monocytogenes.

Image 1. Listeria colonies with beta-hemolysis on blood agar

Discussion

Listeria monocytogenes is an intracellular Gram positive rod that is pathogenic to humans. The organism is a non-spore forming facultative anaerobe which thrives at low temperature and can survive at low pH and high salt concentration, making it an organism of concern in ready to eat- refrigerated food products, including soft cheese, deli meat, and packaged salads. There have been multiple outbreaks of Listeria infection related to certain these food products. The most recent Listeria outbreaks, as per the CDC’s report December 2021, has been linked to Dole packaged salads.

The primary route of infection with Listeria is oral ingestion. The amount of inoculum ingested is responsible for the degree of severity of infection. Most people get noninvasive gastroenteritis, which is self-limited, and recovery typically occurs within a week of infection with supportive treatment. Invasive infection is mostly seen in immunocompromised people, people with hematologic malignancies, chronic illnesses like diabetes, pregnant individuals, and extremes of ages. Infected pregnant woman can transmit the infection to fetus vertically, which can lead to fetal demise. Because of this, pregnant women are advised to avoid foods where Listeria is often found, such as deli meat. The infection rate in US currently is 24 cases per thousand and around 800 cases are reported annually, this number does not include the cases that are unreported which is most likely comprised of the noninvasive gastroenteritis cases.1,2,5

Invasive disease can range from severe form of gastroenteritis to meningoencephalitis. The fetal infection with Listeria is known to result in the most severe form of outcomes ranging from granuloma infantisepticum to fetal demise in utero. Bacteria enter the gastrointestinal tract through the intestinal lining. Upon entry, the bacteria travel intracellularly through the lamina propria into the vascular system and get disseminated throughout the body including the brain and placenta in pregnant woman.1,2,5


Bacterial culture remains the gold standard, of which blood culture is the most sensitive test when it comes to invasive diseases. Blood culture takes about 24 hours to grow the organism. Listeria can be cultured in media containing horse, sheep or rabbit blood. Listeria produces smooth, round, translucent colonies with a narrow zone of beta- hemolysis. Listeria can be morphologically difficult to differentiate from other Gram positive rods. In these cases, simple biochemical tests can help identify the organism. For example, catalase test, esculin test, oxidase test that are typically positive in case of Listeria, while H2S and indole are not produced by the organism and urea and gelatin are not hydrolyzed. Additionally, hanging drop method can help demonstrate the characteristic tumbling motility of the organism. With motility agar, Listeria demonstrates an umbrella motility pattern. Direct identification of organism from a positive blood culture, CSF or tissue specimen by using PCR assays should be performed, when available, and is particularly useful in patient who have undergone antimicrobial therapy. There is microarray based nucleic acid test available that can identify Listeria from blood culture within 3 hours. MALDI-TOF mass spectrometry is an efficient assay for rapid identification of L. monocytogenes once an organism is recovered in culture.2.6

For the treatment of severe listeriosis in at risk populations, ampicillin is commonly used and aminoglycosides can be added for synergy. Trimethoprim/sulfamethoxazole (TMP/SMX) can be given in case of Beta-lactam allergy. Listeria is intrinsically resistant to cephalosporins and therefore these agents should not be used for therapy. In case of suspected treatment failures, antimicrobial susceptibility testing can be performed as per the available CLSI guidelines, which provides susceptibility breakpoints for penicillin, ampicillin, TMP/SMX and meropenem.2,6 Our patient received ampicillin along with gentamicin for her symptoms and recovered well.

References

  1. Radoshevich, L., Cossart, P. Listeria monocytogenes: towards a complete picture of its physiology and pathogenesis. Nat Rev Microbiol 16, 32–46 (2018). https://doi.org/10.1038/nrmicro.2017.126
  2. https://www.cdc.gov/listeria/index.html
  3. Bierne, H. & Cossart, P. When bacteria target the nucleus: the emerging family of nucleomodulins. Cell. Microbiol. 14, 622–633 
  4. Boujemaa-Paterski, R. et al. Listeria protein ActA mimics WASP family proteins: it activates filament barbed end branching by Arp2/3 complex. Biochemistry 40, 11390–11404 (2001)
  5. Lancet Infect Dis. 2017;17(5):510. Epub 2017 Jan 28
  6. Nele Wellinghausen
  7. , 2019. Listeria and Erysipelothrix, Manual of Clinical Microbiology, 12th Edition. ASM Press, Washington, DC. doi: 10.1128/9781683670438.MCM.ch28

-Kritika Prasai, MD. is a PGY-1 Anatomic and Clinical Pathology resident at University of Chicago (NorthShore). 

-Paige M.K. Larkin, PhD, D(ABMM), M(ASCP)CM is the Director of Molecular Microbiology and Associate Director of Clinical Microbiology at NorthShore University HealthSystem in Evanston, IL. Her interests include mycology, mycobacteriology, point-of-care testing, and molecular diagnostics, especially next generation sequencing.

Microbiology Case Study: Worm Seen in Toddler’s Stool

Case History

A worm specimen—as shown in Image 1—was found in a stool sample from a 21 month old, otherwise healthy female.

Image 1. Specimen collect from patient’s stool.

Discussion

The worm specimen in question is Ascaris lumbricoides, the largest of the nematode parasites. Females typically measure 20-35 cm long with straight tails, while males are smaller at 15-31 cm with curved tail.1 A characteristic feature in adults of both sexes are the three “lips” at the anterior end of the body, as shown in Image 2.

Image 2. Close up of the anterior end of an adult A. lumbricoides. Three “lips” are highlighted with a black arrow.

Humans are the definitive host for these roundworm parasites. Infection with these soil-transmitted helminths is quite common, with an estimated 807 million to 1.2 billion people affected.2,3 Children are infected much more frequently than adults.4 Nearly all A. lumbricoides cases occur in tropical and subtropical areas of Asia, sub-Saharan Africa, and the Americas. This infection is rare or absent in developed countries, but sporadic cases may occur in rural regions.3

Individuals affected with adult Ascariasis worms usually show no acute symptoms. However, since these worms are commonly situated in the small intestines, the clinical presentation of a heavy worm burden in children might include stunted growth via malnutrition. In both adults and children, a high worm burden may result in abdominal pain and intestinal obstruction leading to potential perforations. Migrating worms may lead to symptomatic occlusion of the biliary tract, appendicitis, or nasopharyngeal expulsion.3

In the clinical setting and for diagnosis, A. lumbricoides eggs should be found in the feces, juvenile worms in the sputum, and in some cases adults in the feces. For deworming, the recommended treatment are anti-helminthic medications such as albendazole and mebendazole.3 These medications kill the adults, but not the migrating larvae thus repeat treatment might be needed.

References

  1. Centers for Disease Control and Prevention. DPDx – Laboratory Identification of Parasites of Public Health Concern. Internet [updated July 19, 2019]. Available from: https://www.cdc.gov/dpdx/ascariasis/index.html.
  2. Jourdan PM, Lamberton PHL, Fenwick A, Addiss DG. Soil-transmitted helminth infections. Lancet. 2018;391(10117):252-65. Epub 2017/09/09. doi: 10.1016/s0140-6736(17)31930-x. PubMed PMID: 28882382.
  3. Centers for Disease Control and Prevention. Parasites – Ascariasis. Internet [updated November 23, 2020]. Available from: https://www.cdc.gov/parasites/ascariasis/index.html.
  4. Veesenmeyer AF. Important Nematodes in Children. Pediatr Clin North Am. 2022;69(1):129-39. Epub 2021/11/20. doi: 10.1016/j.pcl.2021.08.005. PubMed PMID: 34794670.

-Amelia Lamberty is a Masters Student in the Department of Pathology and Laboratory Medicine at the University of Vermont.

-Christi Wojewoda, MD, is the Director of Clinical Microbiology at the University of Vermont Medical Center and an Associate Professor at the University of Vermont

Microbiology Case Study: A Middle-Aged Female with Fever, Chills, Night Sweats, and Syncope

Case History

A middle-aged female presented to the emergency department after experiencing a fall and loss of consciousness due to syncope. Upon presentation, the patient endorsed an almost four-week history of fevers, chills, abdominal discomfort, night sweats, and dizziness. She also reported poor oral intake and recent unintended weight loss since the onset of her symptoms. When asked, she noted she had returned from a month-long trip to Italy and Ghana two months prior to presentation. She initially presented to an outside hospital with generalized weakness, body aches, and a fever where she was treated with antibiotics for a urinary tract infection. She then presented to a different outside hospital with similar symptoms. There, she confirmed she had not taken malaria prophylaxis and was bitten by mosquitos on her recent trip. Blood was taken for a peripheral blood smear review but no Plasmodium sp. were observed.

At her current presentation, the patient denied a history of seizures but continued to endorse recurrent fevers, malaise, nausea, and vomiting. She was mildly tachycardic, afebrile, and bloodwork revealed normocytic anemia (hemoglobin 10.3), and elevated creatinine. Given the uncertainty surrounding her syncopal episode, the patient was admitted for further workup. After admission, she spiked a fever up to 103°F and the infectious disease service was consulted. As part of her workup, blood was again drawn for Giemsa-stained peripheral blood smears which were read in the microbiology laboratory.

Laboratory Identification

Upon receipt of the patient’s blood, Giemsa-stained thick and thin smears and an immunochromatographic assay for the detection of malarial antigens (BinaxNOW® Malaria, Abbott Laboratories, Abbott Park, IL) were performed. The BinaxNOW® assay was positive for the detection of pan-malarial antigen (T2), but not the histidine-rich protein II antigen specific to P. falciparum (T1). These findings were suggestive of infection with a non-falciparum Plasmodium species (Image 1). Analysis of the Giemsa-stained thin smear revealed several Plasmodium parasites at various stages of development. Importantly, parasites (and particularly ring forms) were only rarely encountered (Image 2, Image 3A). “Basket” (Image 3B) and “Band” (Image 3C) trophozoite forms were observed, as well as schizonts with 6-12 merozoites typical rosette patterns around central pigment (Image 3D). In the context of a positive antigen test, the patient was definitively diagnosed with a Plasmodium malariae infection based on morphology with a calculated parasitemia of less than 0.1%.

Image 1. BinaxNOW® Malaria assay.  This patient’s assay was positive for the common malarial antigen (T2), but the histidine-rich protein II antigen (T1) specific to P. falciparum was not detected.  These results suggest an infection with a non-falciparum Plasmodium species.
Image 2. Developing ring-form trophozoites of P. malariae.  Ring form trophozoites of P. malariae are less-frequently encountered in peripheral smears compared to other Plasmodium species that infect humans.  A) P. malariae rings usually have a single chromatin dot and are generally thicker than that of P. falciparum.  B) As rings develop, the cytoplasm can extend across the cell or can appear with vacuolation leading to “band” or “basket” forms, respectively.
Image 3Gimesa-stained thin smear of erythrocytes infected with P. malariae A)  CellaVision® field with rare infected erythrocytes notated by black arrowheads.  B) “Basket” form trophozoite of P. malariae.  C) “Band” trophozoite of P. malariae.  D) Schizont of P. malariae with 6-12 merozoites surrounding central pigment in a characteristic “rosette”.

Discussion

Plasmodium malariae is one of the five species of Plasmodium (along with P. falciparum, P. vivax, P. ovale and P. knowlesi) which cause human malaria. Infection begins when sporozoites are injected from the salivary glands of the female Anopheles mosquito into the host upon taking a blood meal. Sporozoites migrate to the liver where they infect hepatocytes and develop into schizonts which eventually rupture, releasing infectious merozoites. These merozoites enter the circulation and infect erythrocytes, subsequently developing into immature ring form trophozoites (Image 2A). Ring form trophozoites develop into either mature trophozoites or become gametocytes which can be taken up by another mosquito upon feeding (Image 2B). Mature P. malariae trophozoites adopt unique morphologies not seen with other Plasmodium species including “band” (Image 3B) and “basket” (Image 3C) forms. Mature trophozoites then develop into schizonts (Image 3D) which rupture, releasing 6-12 merozoites which perpetuate the erythrocytic cycle of infection. P. malariae elaborates fewer merozoites than other Plasmodium species which are often arranged in a “rosette” pattern around centrally localized pigment in the schizont (Image 3D).

The P. malariae infectious cycle has several unique hallmarks compared to that of other Plasmodium species. Unlike P. vivax and P. ovale, the P. malariae lifecycle does not include a latent hypnozoite form, and thus is devoid of classical relapse. P. malariae also preferentially infects older erythrocytes, as opposed to P. vivax which prefers younger cells. Additionally, the infected erythrocyte does not enlarge or fimbriate when infected with P. malariae as opposed to P. vivax and P. ovale, respectively. Patterns of erythrocyte infection and lysis lead to elevated parasite burden, characteristic cyclic fevers and anemia. However, the time needed for development from ring trophozoite to rupturing schizont is different among malarial parasites: P. knowlesi exhibits the most rapid development (24-hours), followed by P. falciparum, P. ovale, and P. vivax (48-hours), and then P. malariae (72-hours).  

P. malariae has a global distribution overlapping with P. falciparum. While P. falciparum is the primary species causing reported infection in Ghana, P. malariae infection is encountered less frequently. Associated parasitemia are characteristically lower in P. malariae infections compared to other species due to fewer merozoites produced during infection, an extended 72-hour developmental cycle, and the preference for the infection of older erythrocytes. This can complicate microscopic diagnosis as well as lead to more indolent symptomology. Indeed, patients can often remain asymptomatic for months to years after leaving endemic areas. In this patient’s case, definitive diagnosis was made months following her travel to an endemic region. The patient completed a 5-day course of artemether/lumefantrine with complete resolution of symptoms prior to discharge.

-Clare McCormick-Baw, MD, PhD is an Assistant Professor of Clinical Microbiology at UT Southwestern in Dallas, Texas. She has a passion for teaching about laboratory medicine in general and the best uses of the microbiology lab in particular.

-Andrew Clark, PhD, D(ABMM) is an Assistant Professor at UT Southwestern Medical Center in the Department of Pathology, and Associate Director of the Clements University Hospital microbiology laboratory. He completed a CPEP-accredited postdoctoral fellowship in Medical and Public Health Microbiology at National Institutes of Health, and is interested in antimicrobial susceptibility and anaerobe pathophysiology.

Microbiology Case Study: A Preteen Boy with Cold and Cough

Case History

A preteen boy presented to primary care office with a complaint of flu-like symptoms for the past five weeks. His symptoms improved after 2-3 weeks but noted acute worsening of symptoms in the last two weeks, including sore throat, head congestion, and cough. The physical exam was unremarkable except for nasal congestion, mucosal edema, and some drainage. A chest X-ray was taken, which was normal. Results were negative for a Streptococcal infection, SARS-CoV-2, Bordetella pertussis, and influenza. Bordetella parapertussis was detected by PCR (Image 1).

Image 1. Bordetella pertussis and Bordetella parapertussis PCR. Green = IS481, target gene for B. pertussis. Red = IS1001, target gene for B. parapertussis. Purple = Internal control (IC)

Discussion

Bordetella is a small, non-fermentative, gram negative coccobacilli. The genus Bordetella has 15 species, and B. pertussisB. parapertussis are most commonly found in human infections causing pertussis. B. parapertussis usually cause milder disease, but reports of outbreaks of B. parapertussis have increased in recent literature. The epidemic cycles for pertussis occur at 3–4 years intervals2 and pertussis vaccination does not prevent B. parapertussis infection. B. parapertussis generally occurs in a younger age group than disease caused by B. pertussis.4 Cherry et al. indicated that B. parapertussis infections contribute significantly to the disease burden, which was previously thought to be vaccine failure in children.2

Pertussis is primarily a toxin-mediated disease; the bacteria attach to the cilia of the respiratory epithelial cells and produce toxins that paralyze the cilia and cause inflammation of the respiratory tract, which interferes with the clearing of pulmonary secretions.1 B. pertussis and B. parapertussis are almost identical at the DNA level and produce many similar virulence factors like as filamentous hemagglutinin (FHA), pertactin, tracheal cytotoxin, dermonecrotic toxin, and adenylate cyclase-hemolysin. An essential difference between the two is that B. parapertussis does not secrete pertussis toxin.3,5-9 Despite the high degree of homology shown by the amino acid sequences of the main antigens, the two species differ in respect to several protective epitopes.10

Pertussis (whooping cough) can cause serious illness in babies. Symptoms of pertussis usually develop within 5-10 days of exposure. Early non-specific symptoms, including runny nose, low-grade fever, and occasional cough, can last for 1 to 2 weeks. After 1 to 2 weeks, as the disease progresses, paroxysms occur, which are many, rapid coughs followed by a high-pitched “whoop” sound. Vomiting or exhaustion develops at this stage. Recovery from pertussis is slow, the cough becomes milder and less common, but coughing fits can return with other respiratory infections for many months after the pertussis infection started. The “whoop” is often absent or mild in less severe disease. The illness is generally milder in teens and adults, especially those who have gotten the pertussis vaccine. The cough can be minimal or absent in babies, but they might get apnea, which is most dangerous.1

Bordetella is a fastidious organism as it requires special media, prolonged incubation, timely transport, and rapid plating for recovery of the organism. Regan low and Bordet Gengou are the special media used for culture of B. parapertussis and B. pertussis. Unlike B. pertussis, B. parapertussis can grow on blood and chocolate agar. Colonies may appear like mercury drop and produce beta hemolysis on prolonged incubation. Culture has the highest recovery if a nasopharyngeal swab is collected within two weeks of symptom onset. Sensitivity can be as high as 56% in early disease and decrease over time, while specificity is 100%.1 Serological assay are not clinically validated and do not help differentiate between recent or remote infection or vaccination. PCR is the most sensitive methodology and should be performed from a nasopharyngeal swab taken within three weeks of symptom onset; after the fourth week of cough, the amount of bacterial DNA rapidly diminishes, which increases the risk of obtaining falsely-negative results. PCR-detectable B. pertussis DNA in some pertussis vaccines and the contamination of the clinic environment by those vaccines increases the risk of false-positive PCR. As per CDC guidelines, PCR in asymptomatic persons, asymptomatic close contacts of a confirmed case, and after five days of antibiotic use is unlikely to benefit and is generally not recommended because of the risk of false positivity. In our lab, we use the DiaSorin Simplexa Bordetella direct assay system – RT PCR which targets IS481 and IS1001 for pertussis PCR (other PCR may use different targets). B. pertussis contains ∼238 copies of IS481 and no copies of IS1001, multiple copies of IS481 are responsible for the high sensitivity of PCR and increased risk of false-positive. B. parapertussis has ∼22 copies of IS1001 and no copies of IS481; false-positive identification of IS1001 seems unlikely, as IS1001 is not present in vaccines and its copy numbers are low.2

The recommended antimicrobial agents for treatment or chemoprophylaxis is azithromycin. Antibiotic susceptibility data indicate that the same antibiotics recommended for treating and preventing B. pertussis might help treat and prevent B. parapertussis.11,12 CDC recommends vaccinating young children, preteens, pregnant women, and adults, but pertussis vaccine immunity is short-lived and wanes after 7- 10 years. Immunized children become susceptible after that and can transmit B. pertussis to their very young infant siblings or get B. parapertussis as the vaccine does not protect against it. The average age of patients with B. parapertussis is much younger than those with B. pertussis, and some literature suggest B. parapertussis should be considered when developing new pertussis vaccines.13

References

  1. https://www.cdc.gov/pertussis/index.htmlJames D. Cherry, Brent L. Seaton, Patterns of Bordetella parapertussis Respiratory Illnesses: 2008–2010, Clinical Infectious Diseases, Volume 54, Issue 4, 15 February 2012, Pages 534–537, https://doi-org.proxy.uchicago.edu/10.1093/cid/cir860
  2. Arico B, Rappuoli R. Bordetella parapertussis and Bordetella bronchiseptica contain transcriptionally silent pertussis toxin genes. J Bacteriol.1987;169:2847-2853.
  3. https://www.mayocliniclabs.com/test-catalog/overview/80910#Clinical-and-Interpretive
  4. Blom J., Hansen G. A., and Poulsen F. M.Morphology of cells and hemagglutinogens of Bordetella species: resolution of substructural units in fimbriae of Bordetella pertussis.Infect. Immun.421983308317 CrossrefPubMed.
  5. Cookson B. T. and Goldman W. E.Tracheal cytotoxin: a conserved virulence determinant of all Bordetella species.J. Cell. Biochem.11B1987124
  6. Endoh M., Takezawa T., and Nakase Y.Adenylate cyclase activity of Bordetella organisms. Its production in liquid medium.Microbiol. Immunol.24198095104 PubMed.
  7. Li L. J., Dougan P., Novotny P., and Charles I. G.P70 pertactin, an outer membrane protein from Bordetella parapertussis: cloning, nucleotide sequence and surface expression in Escherichia coli.Mol. Microbiol.51991409417 PubMed.
  8. Mooi F. R., van der Heide H. G. J., TerAvest A. R., Welinder K. G., Livey I., van der Zeisj B. M. A., and Gaastra W.Characterization of fimbrial subunits from Bordetella species.Microb. Pathog.3198718 PubMed.
  9. He Q, Viljanen MK, Arvilommi H, Aittanen B, Mertsola J. Whooping Cough Caused by Bordetella pertussis and Bordetella parapertussis in an Immunized Population. JAMA. 1998;280(7):635–637. doi:10.1001/jama.280.7.635
  10. Hoppe JE, Bryskier A. In vitro susceptibilities of Bordetella pertussis and Bordetella parapertussis to two ketolides (HMR 3004 and HMR 3647), four macrolides (azithromycin, clarithromycin, erythromycin A, and roxithromycin), and two ansamycins (rifampin and rifapentine). Antimicrob Agents Chemother. 1998 Apr;42(4):965-6. doi: 10.1128/AAC.42.4.965. PMID: 9559823; PMCID: PMC105582.
  11. Mortensen JE, Rodgers GL. In vitro activity of gemifloxacin and other antimicrobial agents against isolates of Bordetella pertussis and Bordetella parapertussis. J Antimicrob Chemother. 2000 Apr;45 Suppl 1:47-9. doi: 10.1093/jac/45.suppl_3.47. PMID: 10824032
  12. Karalius VP, Rucinski SL, Mandrekar JN, Patel R. Bordetella parapertussis outbreak in Southeastern Minnesota and the United States, 2014. Medicine (Baltimore). 2017 May;96(20):e6730. doi: 10.1097/MD.0000000000006730. PMID: 28514288; PMCID: PMC5440125.
  13. Karalius VP, Rucinski SL, Mandrekar JN, Patel R. Bordetella parapertussis outbreak in Southeastern Minnesota and the United States, 2014. Medicine (Baltimore). 2017 May;96(20):e6730. doi: 10.1097/MD.0000000000006730. PMID: 28514288; PMCID: PMC5440125.

-Payu Raval, MD is a 1st year anatomic and clinical pathology resident at University of Chicago (NorthShore). Her academic interests include hematology, molecular, and surgical pathology.

-Paige M.K. Larkin, PhD, D(ABMM), M(ASCP)CM is the Director of Molecular Microbiology and Associate Director of Clinical Microbiology at NorthShore University HealthSystem in Evanston, IL. Her interests include mycology, mycobacteriology, point-of-care testing, and molecular diagnostics, especially next generation sequencing.