Microbiology Case Study: A 44 Year Old Male Finds a Tick on His Leg

Case History A 44 year old male pulled this (image 1) off of his leg after dragging brush out of a tree line in Vermont.

Image 1. Ixodes scapularis under a microscope. Characteristic features such as eight black legs, dorsal shield, and dark red color can be appreciated.

Ixodes scapularis

Ixodes scapularis, also known as the blacklegged tick or deer tick, is commonly found in the eastern and northern Midwest regions of the United States as well as southeastern Canada. This species of tick is approximately 3 mm in length. Morphologically, females have a black head and a dorsal shield with a dark red abdomen, while males are entirely black or dark brown. Both sexes have eight black legs and a characteristic anal opening, appearing within a horseshoe-shaped ridge on the ventral lower abdomen. Unlike other tick species, Ixodes scapularis does not have ridges on the edge of the lower abdomen. Ixodes scapularis can live up to 2 years in the wild and die after reproduction.1

Life Cycle, Transmission, and Infection

Ixodes scapularis is a three-host tick with a different host at each stage of development. Their life cycle lasts approximately 2 years, where they undergo 4 distinct developmental/life stages: egg, six-legged larva, eight-legged nymph, and adult. After hatching from the egg, it should have a blood meal at every developmental stage to survive. Ixodes scapularis is known to parasitize and feed from mammals, birds, reptiles, and amphibians, and its best-known host is the white-tailed deer. This species is unable to fly or jump so it usually waits for a host while resting in the tips of grass or shrubs. Depending on the developmental stage, preparation for feeding can take between 10 minutes to 2 hours.2 Once the tick finds a feeding spot on the host, it grasps onto the skin and cuts into the surface inserting its feeding tube, which can have barbs and can secrete a cement-like surface for better attachment. Moreover, the tick can also secrete small amounts of saliva with anesthetic properties to remain undetected during the blood meal. If attached to a sheltered spot, the tick can remain unnoticed for long periods. Ixodes scapularis will attach to its host and suck on the blood for a few days. The lengthy feeding process makes them good at transmitting infection. If the host has a bloodborne infection (e.g., Lyme disease), the tick may ingest the pathogen and become infected. If the tick feeds on a human later, that human can become infected with the same pathogen if it is a prolonged blood meal. However, if the tick is removed quickly (~ 24 hours), the risk of acquiring disease is reduced.2 The longer the tick is attached, the greater the risk of becoming infected. The risk of human infection is greater during the spring and summer.

Ixodes scapularis as a Disease Vector

Babesiosis

The causative agent of babesiosis are Basebesia microti and other Babesia species. These parasites preferentially infect red blood cells. In the United States, most cases are caused by Babesia microti.3 Babesiosis is most frequently reported in the upper midwestern and northeastern regions of the United States, where Babesia microti is endemic. Although this parasite is generally transmitted by Ixodes scapularis, Babesia parasites can also be transmitted via blood transfusions and, in some cases, congenitally. Babesiosis can range from asymptomatic to life-threatening. Some of the common signs and symptoms include fever, chills, sweats, general malaise or fatigue, myalgia, arthralgia, headaches, anorexia, nausea, and dark urine. Less common symptoms include cough, sore throat, emotional lability, depression, photophobia, conjunctival infection.3 Not all infected persons are symptomatic or febrile. Clinical presentation usually manifests within several weeks after exposure, but may develop or recur months after infection. The incubation period for Babesia species parasites is approximately 1-9+ weeks. Laboratory findings associated with babesiosis include decreased hematocrit due to hemolytic anemia, thrombocytopenia, elevated serum creatinine and blood urea nitrogen values, and mildly elevated hepatic transaminase values.3 To diagnose babesiosis in the laboratory, identification of intraerythrocytic Babesia parasites by light-microscopic examination of a blood smear, positive Babesia (or Babesia microti) PCR analysis, or isolation of Babesia parasites from a whole blood specimen by animal inoculation in a reference lab are recommended procedures. Additionally, demonstration of a Babesia-specific antibody titer by indirect fluorescent antibody testing for IgG can be used as supportive laboratory criteria—although it is not enough evidence to support a diagnosis of an active infection.3 Treatment for babesia usually lasts 7-10 days with a combination of two drugs: atovaquone plus azithromycin or clindamycin plus quinine, with the latter being the standard of care for severely ill patients.

Anaplasmosis

Anaplasmosis, formerly known as Human Granulocytic Ehrlichiosis, is caused by Anaplasma phagocytophilum. Anaplasmosis is commonly reported in the upper Midwest and northeastern regions of the United States. The incubation period for Anaplasma phagocytophilum is 5-14 days.3 Some of the common signs and symptoms of anaplasmosis include fever, chills, rigors, severe headaches, malaise, myalgia, gastrointestinal symptoms such as nausea, vomiting, diarrhea, and anorexia, and, in some cases, rash. The general laboratory findings for anaplasmosis during the first week of clinical disease include mild anemia, thrombocytopenia, leukopenia, and mild to moderate elevations in hepatic transaminases.3 Under the microscope, the visualization of morulae in the cytoplasm of granulocytes during examination of blood smears is indicative of diagnosis. However, to definitely determine diagnosis in the laboratory, detection of DNA by PCR of whole blood is recommended during the first week of illness. Additionally, demonstration of a four-fold change in IgG specific antibody titer by indirect immunofluorescence antibody assay in paired serum samples is recommended. The first serum sample should be taken during the first week of illness and the second serum sample should be taken 2-4 weeks after. Moreover, immunohistochemical staining of the organism from the skin, tissue, or bone marrow biopsies is also recommended for diagnosis.3 Anaplasmosis is treated with doxycycline. Treatment should be started once there is a clinical suspicion of disease, as delaying treatment may result in severe illness or in death.

Lyme Disease

The causative agents for Lyme disease include Borrelia burgdorferi and Borrelia mayonii. Lyme disease is most frequently reported in the Upper Midwestern and northeastern regions of the United States with some cases being reported in northern California, Oregon, and Washington. Data from 2015 shows that 95% of Lyme disease cases were reported in the following 14 states: Connecticut, Delaware, Maine, Maryland, Massachusetts, Minnesota, New Hampshire, New Jersey, New York, Pennsylvania, Rhode Island, Vermont, Virginia, and Wisconsin.3 The incubation period for Borrelia parasites is usually 3-30 days.3 Some of the early (3-30 days after a tick bite) signs and symptoms of Lyme disease include fever, chills, headache, fatigue, muscle and joint aches, and swollen lymph nodes may occur with an absence of rash. Erythema migrans is a characteristic rash of Lyme disease and it occurs in 70%-80% of infected people.4 This rash starts at the site of a tick bite after an average of 3-30 days (average is 7 days) and it gradually expands over several days reaching up to 30 cm across.4 As it enlarges, it can result in the characteristic “bulls-eye” appearance; it may feel warm to the touch and it is rarely itchy or painful. Some of the later (days to months after a tick bite) signs and symptoms include severe headache and neck stiffness, additional erythema migrans rashes in other areas of the body, facial palsy, arthritis with severe joint pain and swelling—especially in the knees, intermittent pain in the tendons, muscles, joints, and bones. It may also lead to heart palpitations or Lyme carditis, episodes of dizziness or shortness of breath, inflammation of the brain and spinal cord, nerve pain, and shooting pains, numbness, or tingling of the hands and feet.4 Laboratory diagnosis for Lyme disease includes the demonstration of IgM or IgG antibodies in serum and a two-step testing protocol is highly recommended.5 Moreover, isolation of an organism from a clinical specimen is also recommended. Treatment for Lyme disease includes antibiotics such as doxycycline, cefuroxime axetil, or amoxicillin.

When assessing a patient for any tick-borne diseases, the clinical presentation should be considered alongside the likelihood that the patient has been exposed to an infected Ixodes scapularis tick, or any other tick. Moreover, if a tick is found, engorgement of the tick should be considered when assessing for the possibility of disease transmission.

References

  1. Thevanayagam S. Ixodes scapularis [Internet]. 2012. Available from: https://animaldiversity.org/accounts/Ixodes_scapularis/.
  2. Centers for Disease Control and Prevention. Lifecycle of Blacklegged Ticks [Internet]. 2011 [updated November 15, 2011]. Available from: https://www.cdc.gov/lyme/transmission/blacklegged.html.
  3. Centers for Disease Control and Prevention. Tickborne Diseases of the United States: A Reference Manual for Healthcare Providers [Internet]2018. Available from: https://www.cdc.gov/ticks/tickbornediseases/TickborneDiseases-P.pdf.
  4. Centers for Disease Control and Prevention. Lyme Disease – Signs and Symtoms [Internet]. 2021. Available from: https://www.cdc.gov/lyme/signs_symptoms/index.html.
  5. Mead P, Petersen J, Hinckley A. Updated CDC Recommendation for Serologic Diagnosis of Lyme Disease. MMWR Morb Mortal Wkly Rep. 2019;68(32):703. Epub 2019/08/16. doi: 10.15585/mmwr.mm6832a4. PubMed PMID: 31415492; PubMed Central PMCID: PMCPMC6818702 potential conflicts of interest. No potential conflicts of interest were disclosed.

Amelia Lamberty is a Master’s student in the Pathology Master’s Program.

-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 26 Year Old Female with Diarrhea

Case Description

A 26 year old female with a past medical history of Hemoglobin SC disease (Hb SC) and iron deficiency anemia presented to the emergency department with lower abdominal pain and diarrhea for three days. She began having multiple episodes of watery diarrhea, followed by bloody diarrhea after eating at a restaurant. During this time, she also had fever, chills, body aches, and headache. The patient had been on a course of ceftriaxone and metronidazole started three weeks prior for sore throat, ear infection, and bacterial vaginosis. She completed her metronidazole course prior to the current illness. Abdominal computed tomography revealed splenomegaly and a mildly dilated, fluid-filled appendix without evidence of infectious or inflammatory abnormalities. Hemoglobin on admission was 11.1 mg/dL (Reference Range: 11.2- 15.7 mg/dL) and MCV 62.9 fL (Reference Range: 79.4- 94.8 fL), which is similar to her baseline.

Laboratory Identification

The patient underwent work up for community-acquired diarrhea. Stool cultures grew non-typhoidal Salmonella (Image 1). Blood cultures performed at the time of admission flagged positive with gram negative rods which were also identified as Salmonella species by MALDI-TOF. The organism was susceptible to ampicillin, ceftriaxone, ciprofloxacin, and trimethoprim/sulfamethoxazole. The patient continued on intravenous ceftriaxone and responded to therapy. She was discharged home on oral ciprofloxacin.

Image 1. Salmonella Microbiologic Diagnosis using Xylose Lysine Deoxycholate agar and Triple Sugar Iron slant. A) Non-typhoidal strains of Salmonella are lactose non-fermenting, hydrogen sulfide producing (black colonies) enteric Gram-negative rods on Xylose Lysine Deoxycholate agar (XLD agar). B) Non-typhoidal strains of Salmonella are Alkaline (pink) over Acid (yellow) with the production of copious amounts of hydrogen sulfide on Triple Sugar Iron agar (TSI).

Discussion

Hemoglobin SC disease (Hb SC) is the second most common hemoglobinopathy after Sickle Cell Disease (SCD, Hb SS) globally.1 Hb SC disease occurs when a patient inherits both hemoglobin S and hemoglobin C alleles. Hemoglobin S and C variants are caused by point mutations in the hemoglobin beta- chain, and both variants lead to reduced affinity to the alpha-chain. While hemoglobin C is an abnormal form of hemoglobin that does not cause sickling on its own, when co-inherited with hemoglobin S, the beta chains polymerize, causing red cell sickling when oxygen tension is lowered in the blood.2 Patients develop anemia due to reduced red cell lifespan (27-29 days for Hb SC vs. 15-17 days for Hb SS) and subsequent destruction of red blood cells.3

Complications arise from vascular occlusion and destruction of red blood cells, leading to gallstones, pulmonary infarction, priapism, and/or cerebral infarction. Other complications include avascular necrosis of the femoral head, bone marrow necrosis, renal papillary necrosis, retinopathies, splenomegaly, and recurrent pregnancy loss. Although Hb SC patients often exhibit similar symptomology to sickle cell disease, symptoms are typically milder and present later in childhood.2,3 In comparison to patients with Hb SS, Hb SC patients have milder anemia, less frequent sickle cells, and less severe hemolysis. While Hb SC patients have fewer sickling episodes compared to Hb SS patients, Hb SC patients have more severe retinopathy and splenomegaly. It is also important to note that the enlargement of the spleen is often caused by red blood cell sequestration and the optimal function of the spleen is significantly reduced (functional hyposplenia), which can lead to increased risk of infection from encapsulated bacteria.

Diagnosis of Hb SC disease is typically made by performing hemoglobin electrophoresis (Image 2). Hemoglobin electrophoresis separates the differing varieties of hemoglobin by size and electrical charge. Capillary electrophoresis separates hemoglobin variants based on the “zone” of detection where each variant hemoglobin appears based on a reference pattern. Normal hemoglobin (A, F, A2) is easily discriminated from variant hemoglobins (S, C, E, D), and quantification allows for detection of beta-thalassemia (increased A2 fraction). While useful as a screening tool, the hemoglobin variants identified in the “zones” are not specific. For example, Hb C and Hb Constant Spring share a zone, and Hb A2 shares a zone with Hb O- Arab. Variants detected by capillary electrophoresis are confirmed by a second method, and in this case Hb SC was confirmed by acid agarose gel (Sebia Hydrogel). When subjected to acid gel electrophoresis, Hb C and Hb S migrate in separate bands, while Hb A, A2, D, and E comigrate in the “A” band, and the “F” band may contain F in addition to the glycated fraction of normal adult Hb A. Patients with Hb SC disease will have variants detected in the S and C zones in capillary electrophoresis and lack signal in the A zone.4

Image 2. Laboratory Diagnosis of Hb SC disease includes hemoglobin electrophoresis and peripheral blood smear review. A) Hemoglobin capillary electrophoresis (pH 9.4) separates F, S, C, A2, A (Sebia, Capillarys 2 Flex Piercing). B) Acid agarose gel (pH 6.0-6.2) separates hemoglobins F, A, S, and C (Sebia, Hydragel Acid QC lane).  C) Peripheral blood smear morphology showing characteristic Hb SC forms including target cells, boat shaped cells (single arrow), red cell with crystals (double arrow), and hemighost cells (triple arrow).

Examination of the peripheral blood smear from a patient with Hb SC disease (Image 2C) reveals frequent target cells, boat-shaped cells (taco shaped), and only rarely contains classic sickle cells. Hemoglobin C crystals can be seen, both free floating and inside red cells, a feature of CC and SC disease but not seen in SS disease. Hemi-ghost cells and cells with irregular membrane contractions are also more frequent in Hb SC disease. In contrast, sickle cells are rarely observed in peripheral smears from Hb SC patients.

Salmonellaeare flagellated gram negative bacilli that are members of the Enterobacterales. Salmonellosis is typically foodborne in nature and presents as a self-limiting acute gastroenteritis.5,6 However, these organisms can invade beyond the gastrointestinal tract resulting in bacteremia.6 This case presents Salmonella as a cause of bacteremia in a patient with Hb SC disease following a bout of gastroenteritis. Although there is a well-known association between SCD and invasive infections with Salmonella, the incidence of Salmonella infection in patients with Hb SC disease has not been well studied. Patients with SCD, particularly those in Africa, are at risk for developing invasive disease caused by non-typhoidal Salmonella, including osteomyelitis, meningitis, and bacteremia. It has been hypothesized that disruptions in the gut microbiome and increased permeability of enterocytes makes SCD patients more prone to invasive Salmonella infections.6 Furthermore, the compromised function of the spleen in both patients with SCD and Hb SC disease increases the risk of disseminated infection by encapsulated bacteria and Gram negative rods. The spleen plays an important housekeeping role removing old or damaged erythrocytes, but also has an important immunological function housing memory B cells, producing antibodies and macrophages that phagocytize circulating bacteria, particulates or other debris and then present the antigens to other immunological cells in the spleen.7 Although sepsis caused by Salmonella is an occasional progression of gastroenteritis, this patient’s Hb SC disease likely increased the likelihood of bacteremia because of her functional asplenia.

References

  1. Weatherall DJ. The inherited diseases of hemoglobin are an emerging global health burden. Blood. 2010;115(22):4331–6.
  2. Tim R. Randolph,24 – Hemoglobinopathies (structural defects in hemoglobin),Editor(s): Elaine M. Keohane, Catherine N. Otto, Jeanine M. Walenga,Rodak’s Hematology (Sixth Edition), Elsevier, 2020, Pages 394-423, ISBN 9780323530453, https://doi.org/10.1016/B978-0-323-53045-3.00033-7.
  3. (https://www.sciencedirect.com/science/article/pii/B9780323530453000337)
  4. Nathan, D. G., Orkin, S. H., & Oski, F. A. (2015). Sickle Cell Disease. In Nathan and Oski’s hematology and oncology of infancy and childhood (8th ed., pp. 675-714). Philadelphia, PA: Elsevier. Retrieved from https://www.clinicalkey.com/#!/content/book/3-s2.0-B9781455754144000206y.com/#!/content/book/3-s2.0-B9781455754144000206. Accessed 2022
  5. Bain, BJ. (2020) Haemoglobinopathy Diagnosis, Third Edition. Hoboken: John Wiley and Sons, Ltd
  6. Kurtz, J. R., Goggins, J. A., & McLachlan, J. B. (2017). Salmonella infection: Interplay between the bacteria and host immune system. Immunology letters190, 42–50. https://doi.org/10.1016/j.imlet.2017.07.006
  7. Lim, S.H., Methé, B.A., Knoll, B.M. et al. Invasive non-typhoidal Salmonella in sickle cell disease in Africa: is increased gut permeability the missing link?. J Transl Med 16, 239 (2018). https://doi.org/10.1186/s12967-018-1622-4
  8. Leone G, Pizzigallo E. Bacterial Infections Following Splenectomy for Malignant and Nonmalignant Hematologic Diseases. Mediterr J Hematol

-John Stack is a first year AP/CP resident at UT Southwestern Medical Center.

-Marisa Juntilla is an Assistant Professor in the Department of Pathology at UT Southwestern Medical Center. Dr. Juntilla is a board certified Clinical Pathologist and is certified in the subspecialty of Hematopathology.

-Dominick Cavuoti is a Professor in the Department of Pathology at UT Southwestern Medical Center. Dr. Cavuoti is a board certified AP/CP who is a practicing Clinical Microbiologist, Infectious Disease pathologist and Cytopathologist.

-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.

-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.

Microbiology Case Study: An Adult Presents with Hand Wound Following a Dog Bite

Case Presentation

An adult presented to the emergency department with a finger infection persisting for the past 14 days after being bitten by her dog. The finger was swollen, tender and red but the patient denied fever, chills, or purulent drainage. The patient was previously given 10 days of doxycycline and amoxicillin-clavulanic acid without any improvement. The patient underwent incision and drainage and the specimen was sent for aerobic culture and Gram stain. No organisms or WBCs were seen on the Gram stain. On day 3 of incubation, a yellow colony was observed on the chocolate agar. The colony was streaked out onto another chocolate plate for subculture (Image 1). MALDI-TOF identified this organism as Neisseria animoralis.

Image 1. Subculture of Neisseria animoralis.

Discussion

Neisseria animoralis and Neisseria zoodegmatis are primarily zoonotic organisms found as normal oral flora of cats and dogs. Both can cause wound infections in humans following animal bites. However, these organisms are under recognized animal bite pathogens, often leading to their identifications being dismissed as contaminants. While there are limited published studies on this organism, it is important to recognize its role in wound infections, as in our case. Due to lack of awareness and reduced recovery in culture, case studies have shown correlations with this organism and poor healing and chronic wound infections.

On Gram stain, N. animoralis appears as a Gram negative coccoid rod. In culture, N. animoralis is a slow growing organism that produces yellow or white colonies that are shiny and smooth. N. animaloris produces acid from glucose, but not lactose, sucrose, or maltose. MALDI-TOF is most commonly used for identification.

Limited N. animoralis treatment data are available currently. Most animal bite-related infections are polymicrobial in nature and thus, antibiotic treatment is broad spectrum to cover the most common aerobic and anaerobic organisms.

Resources

  • Johannes Elias, Matthias Frosch, and Ulrich Vogel, 2019. Neisseria, In: Carroll KC, Pfaller MA Manual of Clinical Microbiology, 12th Edition. ASM Press, Washington, DC. doi: 10.1128/9781683670438.MCM.ch36
  • Heydecke A, Andersson B, Holmdahl T, Melhus A. Human wound infections caused by Neisseria animaloris and Neisseria zoodegmatis, former CDC Group EF-4a and EF-4b. Infect Ecol Epidemiol. 2013;3:10.3402/iee.v3i0.20312. Published 2013 Aug 2. doi:10.3402/iee.v3i0.20312
  • Kathryn C. Helmig, Mark S. Anderson, Thomas F. Byrd, Camille Aubin-Lemay, Moheb S. Moneim, A Rare Case of Neisseria animaloris Hand Infection and Associated Nonhealing Wound, Journal of Hand Surgery Global Online, Volume 2, Issue 2, 2020, Pages 113-115, ISSN 2589-5141 https://doi.org/10.1016/j.jhsg.2020.01.003.
  • Merlino J, Gray T, Beresford R, Baskar SR, Gottlieb T, Birdsall J. Wound infection caused by Neisseria zoodegmatis, a zoonotic pathogen: a case report. Access Microbiol. 2021;3(3):000196. Published 2021 Feb 10. doi:10.1099/acmi.0.000196

-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: 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.