An Adult Patient Presents with Mild Penile Irritation and Discharge

Patient History

An adult male presented to the primary care office with mild penile irritation and discharge without fever, dysuria, or other lesions. He is sexually active and reported recent unprotected sex with multiple partners. He is on pre-exposure prophylaxis for HIV and tested non-reactive for HIV, HCV, and syphilis antibodies. Chlamydia and gonorrhea were detected in his urine, rectal, and throat specimens by PCR. The lab paged the director to review and verify the results. Is it possible to be positive for both chlamydia and gonorrhea?

Discussion

In the United States, chlamydia and gonorrhea are the most commonly reported sexually transmitted bacterial infections. While most cases of chlamydia and gonorrhea are sexually transmitted, neonates can become infected by perinatal transmission.1,2,3 To prevent long-term complications in women, all sexually active women aged <25 years and older women with increased risk of infection should get tested annually for chlamydia and gonorrhea. All pregnant women <25 years old or are considered high risk should be screened at the first prenatal visit and in the third trimester or at the time of delivery for both organisms. CDC recommends screening genital and extragenital sites at least annually for all sexually active MSM at risk for infection.4

Chlamydia trachomatis (C. trachomatis) is a gramvnegative, obligate, aerobic, coccoid or rod shape bacteria that does not grow in routine culture. C. trachomatis cannot synthesize ATP and humans are the only known natural host for C. trachomatis.4 Neisseria gonorrhoeae (N. gonorrhoeae) is a Gram-negative, facultatively intracellular, obligate aerobe diplococci. While this organism can be grown in culture, sensitivity is lower compared to routine molecular methods. Co-infection is common, with an estimated 10–40% of patients with gonorrhea also infected with chlamydia, and the data also suggested an interplay between these two pathogens. 5,6,7 Patients with chlamydia and gonorrhea co-infection can have increased gonococcal bacterial load, which might facilitate gonorrhea transmission compared with a single infection. Chlamydia can evade the host immune response by preventing neutrophil extracellular traps (NETs) production, which can help gonorrhea to establish intracellular infection.8 Studies in mice suggest that C. trachomatis induces changes in the genital tract immune environment, making it a more permissive environment for N. gonorrhoeae.9

Appropriate specimens include self- or clinician-collected vaginal swab, endocervical swab, urethral swab, and first catch urine. For chlamydial and gonococcal infection diagnosis, CDC recommends testing by nucleic acid amplification tests (NAATs). NAATs are more sensitive and specific compared to other methods. FDA has approved NAATs for urogenital specimens and only particular platforms are approved for rectal and oropharyngeal specimens. C. trachomatis does not grow in routine culture and diagnosis at this time relies solely on NAAT. For N. gonorrhoeae, culture and antibiotic susceptibility should be evaluated in case of suspected treatment failure. Our lab uses Abbott Real-time CT/NG assay, which is currently FDA approved for testing urogenital specimens only.

CDC recommends treating chlamydia with a seven-day course of doxycycline with sexual abstinence until treatment completion/resolution of symptoms. Azithromycin or levofloxacin can be used as alternatives. For gonorrhea, a single ceftriaxone intramuscular injection is recommended, and gentamicin with azithromycin can be used in case of cephalosporin allergy. Unfortunately, for pharyngeal gonorrhea, there is no reliable alternative available for ceftriaxone allergy. Sexual partner evaluation, testing, and presumptive treatment are recommended, along with patient treatment.10 In cases where the chlamydial infection has not been ruled out, patients should also receive anti-chlamydial therapy. A test-of-cure (follow-up testing) for gonorrhea is required in throat infections only after 14 days of the treatment.10

References:

  1. Kreisel KM, Spicknall IH, Gargano JW, Lewis FM, Lewis RM, Markowitz LE, Roberts H, Satcher Johnson A, Song R, St. Cyr SB, Weston EJ, Torrone EA, Weinstock HS. Sexually transmitted infections among US women and men: Prevalence and incidence estimates, 2018. Sex Transm Dis 2021; in press.
  2. CDC. Sexually Transmitted Disease Surveillance, 2020. Atlanta, GA: Department of Health and Human Services; April 2022.
  3. https://www.cdc.gov/std/chlamydia/stdfact-chlamydia-detailed.
  4. http://dx.doi.org/10.15585/mmwr.mm6950a6external icon

-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.

Triaging Times

As a clinical instructor and lead cytologist at my institution, I like to remind our newer cytologists and cytology students of the importance of being prepared for FNA biopsies so they develop good habits or best practices as they become more experienced. This level of preparation helps to create a culture of ongoing learning and improvement, which is necessary for the laboratory. In my experience, I’ve met some cytologists who prefer to go into a case blind, with the mindset that knowing the patient’s clinical history in advance muddies their knowledge, skills, and abilities, limiting their mindset by excluding the possibility of other diagnoses. While diving into the unknown might seem exciting, it is also a hindrance and could result in errors, especially when the clinical history helps us triage the patient’s sample. For example, knowing that the patient has a history of lymphoma or that the presentation state includes bulky lymphadenopathy prompts us to collect additional needle passes to send for flow cytometry analysis. Another concern is not knowing whether the patient has a history of breast, gastric, or esophageal cancer, and consequently processing the specimen routinely, which may result in an extended cold ischemic time. This delay in fixation along with insufficient formalin fixation can yield false negatives on ER/PR IHC in breast cancers and HER2 FISH in breast, gastric, and esophageal cancers, which could restrict the use of hormone therapies, such as tamoxifen and aromatase inhibitors for hormone receptor-positive (HR+) cancers, or trastuzumab for HER2+ cancers. I cannot overemphasize the importance of familiarizing yourself with clinical history and communicating case specifics while you act as a mediator between clinician and pathologist.

Whether the clinical history impacts the pre-analytical phase, such as specimen collection (limiting cold ischemic time or collecting additional needle passes for ancillary studies) or the analytical phase, as such processing (formalin fixation) and diagnosis (selecting an appropriate immunoprofile), we must remain vigilant and proactive in laboratory medicine. In this case, knowing the patient’s clinical history was of the utmost significance as it helped to reduce the number of immunostains and ancillary studies necessary to make the diagnosis. Using morphologic criteria in tandem with the patient’s clinical history narrowed the differential diagnoses to just two possible types of cancer, presented below.

A 59 year old male patient presented to the emergency room after an automobile accident. On imaging, the X-ray and CT scan identified a left humerus mass and fracture, and bloodwork was performed. His medical record was sparse and uneventful with no recent visits or encounters. To build a more comprehensive wellness profile and prepare for surgery, he was also offered a one-time screening for Hepatitis C, as an adult who was born between 1945 and 1965.

The left humerus mass was biopsied via CT-scan guidance and two passes were obtained. The Diff-Quik stained smears demonstrate large polygonal cells, some with abundant, granular cytoplasm and some isolated cells with naked nuclei. Vessels also appear to traverse some of the cell groups.

Images 1-2: Bone, Humerus, Left, CT-guided FNA. Diff-Quik-stained smears.

The Pap-stained smears also demonstrate polygonal cells with granular cytoplasm, nuclei with coarse chromatin, and prominent nucleoli. An interesting feature frequently identified in this case is the intranuclear inclusions, and in hindsight, a focus on these may have further reduced the number of immunostains performed.

Images 3-5: Bone, Humerus, Left, CT-guided FNA. Pap-stained smears.

The H&E-stained cell block sections show trabeculae with endothelial wrapping around the cell cords. While renal cell carcinoma was listed as a differential diagnosis due to its telltale oncocytic cytoplasm and vascularity, hepatocellular carcinoma was favored.

Images 6-7: Bone, Humerus, Left, CT-guided FNA. H&E sections (6: 100x, 7: 400x).

Immunostains were performed using proper positive and negative controls on the cell block sections, and the tumor cells show positive staining for Arginase, cam5.2, and Hepar1, while negative staining for CK7 and PAX8 (not shown).

Images 8-10: Bone, Humerus, Left, CT-guided FNA. Cell block section immunohistochemistry. 8: Arginase-positive; 9: cam5.2-positive; 10: Hepar1-positive.

Fortunately, before ordering immunostains, both our cytologist and pathologist working on the case peered into the patient’s medical record and noticed that he had recent bloodwork which demonstrated a positive Hepatitis C screening. This diagnosis was as recent as the identification of his humerus mass. Had it not been for his car accident, I can’t imagine how long he would have gone undiagnosed for both hepatitis and metastatic hepatocellular carcinoma. Incidental findings save lives, folks.

Granted, in settings of unknown primaries with widespread metastatic disease, such as carcinomatosis, an extensive workup is almost always inevitable. Narrowing down possible etiology based on information such as gender, age, and environmental or occupational exposure can help, but that doesn’t always yield a definitive answer as time- or cost-effectively as possible. In this case, that one clue of untreated Hepatitis C was all the cytopathology team needed. A rarity, sure, but as we are asked to do more personalized tests with less material, think of the patient’s specimen as a puzzle and keep your eye out for a clue both under the microscope and behind the computer. You never know what you might find that reduces errors and unnecessary testing while efficiently leading to a definitive diagnosis.

-Taryn Waraksa-Deutsch, MS, SCT(ASCP)CM, CT(IAC), has worked as a cytotechnologist at Fox Chase Cancer Center, in Philadelphia, Pennsylvania, since earning her master’s degree from Thomas Jefferson University in 2014. She is an ASCP board-certified Specialist in Cytotechnology with an additional certification by the International Academy of Cytology (IAC). She is also a 2020 ASCP 40 Under Forty Honoree.

Microbiology Case Study: Severely Immunocompromised Female with Respiratory Failure

Case History

A 50 year old female with a complex medical history consisting of lymphoma, diabetes mellitus (type II), sarcoidosis, congestive heart failure, chronic renal failure (stage 3), and pancytopenia  presented to the emergency department with shortness of breath, cough, fever. She was found to be positive for SARS-CoV-2 and was transferred to the ICU due to hypoxic respiratory failure. She was treated for sepsis and respiratory failure, but her status continued to decline. The patient had multiple admissions due to COVID-19 in the past, received remdesivir and was on corticosteroid therapy due to the interstitial lung disease from last year. Initial evaluation included complete blood count which revealed anemia (hemoglobin=8.7 mg/dl), leukocytosis (WBC = 21,900/mcl), lymphopenia (910/mcl) and thrombocytopenia (Plt = 27000/mcl). The patient was treated with broad antibiotics and additional steroids. Additional tests revealed hyperproteinemia and hypoalbuminemia. Chest x-ray showed worsening infiltrates in lungs and chest CT scan revealed left apical hydropneumothorax, loculated left pleural effusion, pneumomediastinum, and chest wall subcutaneous emphysema. Lung biopsy revealed necrosis. Histopathology examination revealed broad, branching hyphae with sporulation in lung tissue biopsy and bronchoalveolar lavage. Respiratory cultures of lung biopsy and BAL grew rapidly and lactophenol cotton blue tape preps showed broad hyphae with round sporangium and rhizoids between the stolons. The patient was diagnosed with mucormycosis, infection with Rhizomucor, and was treated with Amphotericin B. Surgical debridement of the tissue was not possible due to her declining condition. She passed away after 5 days.

Figure 1. H&E stain of the lung biopsy (top, left) and Papanicolaou stain of bronchoalveolar lavage (top, right) revealed broad, ribbon-like, right-angle branching hyphae (visible in lung biopsy) with sporulation (credits to Dr. Elham Arbzadeh, George Washington University School of Medicine and Health Sciences). Rapid growth was observed from the respiratory cultures of the tissue biopsy by day 2 (bottom, left) where lactophenol cotton blue tape preps showed broad hyphae with sporangium (bottom, right) and intermodal rhizoids (not shown in this image).

Discussion

The term mucormycoses refers to infections caused by the Zygomycetes which is further separated into Mucorales and Entomophthorales. Some of the members of Mucorales are Rhizopus spp., Mucor spp., Lichtheimia (Absidia) spp., Syncephalastrum spp., and Rhizomucor spp.1,2 These organisms live in soil, dung, and vegetative matter. Infection is usually acquired by inhalation/ingestion of their spores or direct inoculation and contamination of wounds. The mold can invade the walls of the blood vessels causing angioinvasion and often results in dissemination of mycotic thrombi and development of systemic infection. Zygomycetes are most commonly known for causing rhinocerebral, pulmonary, cutaneous, and disseminated disease. Infections with Zygomycetes most commonly occur as opportunistic infections in immunocompromised hosts. Risk factors include diabetes, those with acidosis, neutropenia, and sustained immunosuppression such as after transplantation.

Zygomycetes grow very fast (within 48 to 72 hrs.) and is often called a “lid lifter”. The colonies have a wooly mycelium and can be described as cotton candy-like. Lactophenol tape preps of the mold would reveal broad hyphae, aseptate or pauciseptate, ribbon-like hyphae with irregular width. At the tip of the sporangiophore, there is a sack-like structure called a sporangia with contains all the spores. Fungal elements and hyphae seen on tissue biopsies from patients with mucormycosis typically have near right angle branching (usually >40o) broad, non-septate hyphae. In contrary, those with aspergillosis show acute angle branching (usually <45o) with narrow, septate hyphae.3  

Genus-level identification can be achieved by microscopic morphology. Rhizomucor is an intermediate between Rhizopus and Mucor. Rhizoids found in Rhizomucor are few in number and are located on stolons, between the sporangiophores, as opposed to Rhizopus where the rhizoids are often seen directly at the nodes and Mucor which does not produce rhizoids. Sporangia (40-80 µm in diameter) are brown in color and round in shape. Apophysis is absent, which allows for differentiation from Lichtheimia (Absidia) where apophysis can be seen.4 The genus Rhizomucor includes three species: Rhizomucor pusillusRhizomucor miehei, and Rhizomucor tauricus.5

Treatment of mucormycosis consists of antifungal and surgical therapy. Amphotericin B is the most commonly used antifungal agent. Liposomal amphotericin B has also been successfully used in some cases with zygomycosis due to Rhizomucor.6  Early diagnosis and treatment are crucial and mortality rate is high.7  Of note, Zygomycetes are intrinsically resistant to voriconazole.

References

  1. Rippon J W. Medical mycology. The pathogenic fungi and the pathogenic actinomycetes. Philadelphia, Pa: Saunders; 1974. Mucormycosis; pp. 430–447. 
  2. Scholer H J, Müller E. Beziehungen zwischen biochemischer Leistung und Morphologie bei Pilzen aus der Familie der Mucoraceen. Pathol Microbiol. 1966;29:730–741.
  3. Mohindra S., Mohindra S., Gupta, R., Bakshi, J., Gupta, S. K. Rhinocerebral mucormycosis: the disease spectrum in 27 patients. Mycoses. doi: 10.1111/j.1439-0507.2007.01364.x.
  4. de Hoog, G. S., J. Guarro, J. Gene, and M. J. Figueras. 2000. Atlas of Clinical Fungi, 2nd ed, vol. 1. Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands)
  5. Schipper M A A. On the genera Rhizomucor and Parasitella. Stud Mycol. 1978;17:53–71. 
  6. Bjorkholm, M., G. Runarsson, F. Celsing, M. Kalin, B. Petrini, and P. Engervall. 2001. Liposomal amphotericin B and surgery in the successful treatment of invasive pulmonary mucormycosis in a patient with acute T- lymphoblastic leukemia. Scand J Infec Dis. 33:316-319.
  7. Ribes, J. A., C. L. Vanover-Sams, and D. J. Baker. 2000. Zygomycetes in human disease. Clin Microbiol Rev. 13:236-301.

-Maryam Mehdipour Dalivand, MD is a Pathology Resident (PGY-1) at The George Washington University Hospital. She is pursuing AP/CP training.

-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.

Tick Identification: Why Do We Do It and What Does It Tell Us?

During the warmer months here in the Midwest, ticks are abundant and our microbiology lab receives several tick submissions per day for identification. When possible, we provide species level identification as well as sex for any tick submitted. While this is common practice in most microbiology laboratories, our molecular laboratory accidently received a tick specimen and, in the process of routing it to the microbiology lab, was curious as to why the tick identification matters—what does that tell us clinically? This led to an impromptu plate rounds with both labs and prompted me to write this post.


How do we determine tick identity?

A tick is submitted in a cup and sent to the laboratory. Ideally the tick would be submitted whole without missing appendages or damaged in any way. The tick is placed in ethanol to kill the organism and to allow for examination under a microscope. The mouth parts, scutum, and festoons are examined for defining features. Thorough examination is challenging when the tick arrives damaged or only partially intact.

Why do we provide tick identification?

Certain ticks carry specific pathogens. For instance, Amblyomma americanum (lone star tick) can transmit ehrlichiosis, Francisella tularensis, Heartland virus, Bourbon virus, and Southern tick-associated rash illness, while Ixodes scapularis can transmit Borrelia burgdorferi & Borrelia mayonii (both are causative agents of Lyme disease), Anaplasma phagocytophilum, and Erhlicia muris as well as Powassan virus. Knowing which tick that the patient was bitten by can allow providers to understand what potential pathogens they may or may not have been exposed to. If Amblyomma americanum is submitted, for example, that tick does not carry Borrelia burgdorferi. However, it is important to note that the majority of patients who develop tick-borne illness have no recollection of a tick bite! So while one tick may be discovered and sent to the lab, the patient could still have been unknowingly bitten by a different tick, which could carry other pathogens. When a patient exhibits clinical symptoms that are consistent with a tick-borne disease, such as Lyme Disease, the patient should be tested for that disease regardless of their tick history.

The patient has an Ixodes tick! They are worried about Lyme Disease. Should we send the tick out for molecular testing?

We discourage the use of molecular testing on the ticks themselves because ticks carry a variety of pathogens and there is a high likelihood of carrying a particular pathogen in a high prevalence area. For Ixodes ticks in Lyme Disease endemic areas, 15-70% of ticks may carry the causative agent, Borrelia burgdorferi. However, just because a tick carries a particular pathogen, it does not mean that the patient is now infected. This can lead to unnecessary treatment and misdiagnosis. Moreover, ticks must feed for a certain amount of time before pathogens can be transmitted. For example, Ixodes ticks must typically feed for more than 24 hours before it can transmit Lyme Disease or other pathogens.

Image 1. A male Dermacentor variabilis (also known as the American dog tick) submitted by one of our patients.

In summary, tick identification can provide a glimpse into what the patient was potentially exposed to and if symptoms do arise days to weeks later, the tick identification may offer additional clues. However, just because a person was bitten by a tick does not mean that they are infected. Identification is just a piece of the puzzle!

References

-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 47 Year Old with Eye Pain and Redness

Case history

A 47 year old male with an extensive ocular history including laser assisted in situ keratomileusis (LASIK), multiple ocular traumas with repair, and myopic degeneration with neovascularization for which he was prescribed hard and soft contact lenses presented for bilateral eye redness, watering, and stinging pain. Recently, he had forgotten his soft contacts and wore his hard lenses to his job in construction which he reported doing once every 1-2 months. Ocular exam revealed only his usual chronic changes. His symptoms improved with moxifloxacin eyedrops, but never fully resolved. A month later he returned with what was initially assessed as diffuse corneal edema and conjunctival injection in his left eye, but no ring infiltrate or epithelial defect. Two days later, a large epithelial defect with surrounding ring infiltrate and hypopyon (settling of white blood cells at the base of the anterior chamber) developed in his left eye. Confocal microscopy showed findings concerning for Acanthamoeba infection and the contact lenses and case were sent for culture. Environmental organisms including Klebsiella varicola, Chryseobacterium gleum, and Pseudomonas fluorescens were recovered. In addition, cultures for Acanthamoeba sp., where sample is overlaid on a lawn of E. coli grown on a non-nutrient agar plate, were sent to a reference laboratory.

The patient was treated with Brolene, polyhexamide biguanide (PHMB), and chlorhexidine for Acanthamoeba as well as with antibacterial agents. Three months later his LASIK flap failed and was removed and sent for cultures and pathology which both grew Acanthamoeba sp.(Image 1). He continued treatment for another two months, but the corneal defect expanded. He underwent a therapeutic penetrating keratoplasty, and the explant cornea was sent for pathology. Sections showed acute and chronic inflammation of the corneal epithelium and stroma with rare cysts of Acanthamoeba with atypical morphology possibly representing treatment effect or nonviable organisms (Image 2). The patient continued treatment for another month afterward with resolution of symptoms.

Image 1. Representative photomicrographs of cornea with multiple Acanthamoeba cyst forms at differing stages of development (H&E, 400x magnification) and trophozoite with associated acute inflammation (inset, 500x magnification, oil immersion).
Image 2. Photomicrograph of this patient’s explanted LASIK flap. A) Low power magnification demonstrating acute and chronic inflammation in a background of degrading corneal tissue. An empty cyst is highlighted by the arrowhead (H&E, 100x magnification). B and C) High power magnification of likely nonviable cysts indicated by the arrowheads (H&E, 400x magnification).

Discussion

Acanthamoeba sp. are free-living amoebae found ubiquitously in the environment including in water, soil, dust, and air conditioning ducts.1 Over 20 species of Acanthamoeba have been identified, with eight known to cause human disease. A. castellani and A. polyphaga are the most common species identified from clinical infections.2 Acanthamoeba sp. are a primary reservoir of Legionella pneumophilia and can serve as vectors for other bacterial infections.3 These organisms may colonize the nasal passages of normal hosts.4 Acanthamoebal infections have varied clinical presentations depending on the route of transmission, organ(s) infected, and immune status of the host. These include amebic keratitis, granulomatous amebic encephalitis, and disseminated disease.3 Of these, Acanthamoeba keratitis (AK) is the most frequently encountered clinically.

AK can occur when the organisms are inoculated into corneal micro-abrasions, most often from contaminated hard contact lenses rinsed with tap water. AK represents 5% of all cases of contact-lens-associated keratitis, and 70-85% of AK cases are associated with contact lens use.1 Diagnosis of AK is heavily dependent on a high index of suspicion as AK presents with nonspecific ocular symptomology including blurred vision, photophobia, inflammation, and eye pain. A corneal ring infiltrate is characteristic, but only present in 50% of cases.1 Although historically culture is the gold standard for diagnosis, advanced technologies like confocal microscopy and PCR have greatly improved sensitivity and time to diagnosis.5 Cultures are usually grown on agar plates coated with gram negative bacilli such as E. coli.2 If Acanthamoeba are present, trails of bacterial clearing can usually be seen within days but may take up to several weeks.2 They have dormant cyst and active trophozoite forms. Microscopically they appear as round heterogeneous bodies with a distinct nucleus and surrounded by ruffled membrane and are 15-35 μm in length.3 PCR, given its analytical sensitivity, specificity and turn around time, is the more common method of diagnosis of AK and has replaced many instances of culture today.

AK has a poor prognosis and is potentially sight threatening. Factors contributing to disease severity include delayed diagnosis, pathogenic factors, and lack of effective medical management.1 Nearly 40% of patients fail initial therapy.1 Factors that contribute to Acanthamoeba pathogenicity include production of enzymes including elastases and proteases, adhesion molecules, and physiologic tolerance to different temperatures, osmolarities, and pH.6 The cyst stage confers resilience to many therapies which is compounded by poor tissue penetration of the antimicrobial agents often used in therapy.6 Repeated exposure to therapeutic antimicrobials can also lead to the development of resistance.6 In our patient’s case, treatment was successful following the LASIK flap removal, facilitating increased drug penetration and supported by pathologic findings of treatment effect in the explanted cornea.

References

  1. Somani SN, Ronquillo Y, Moshirfar M. Acanthamoeba Keratitis. 2021 Aug 11. In: StatPearls [Internet]. Treasure Island (FL): StatPearls Publishing; 2021 Jan–. PMID: 31751053.
  2. Maycock NJ, Jayaswal R. Update on Acanthamoeba Keratitis: Diagnosis, Treatment, and Outcomes. Cornea. 2016 May;35(5):713-20. doi: 10.1097/ICO.0000000000000804. PMID: 26989955.
  3. Marciano-Cabral F, Cabral G. Acanthamoeba sp. as agents of disease in humans. Clin Microbiol Rev. 2003 Apr;16(2):273-307. doi: 10.1128/CMR.16.2.273-307.2003. PMID: 12692099; PMCID: PMC153146.
  4. Clarke B, Sinha A, Parmar DN, Sykakis E. Advances in the diagnosis and treatment of Acanthamoeba keratitis. J Ophthalmol. 2012;2012:484892. doi: 10.1155/2012/484892. PMID: 23304449; PMCID: PMC3529450.
  5. Hoffman, J.J., Dart, J.K.G., De, S.K. et al. Comparison of culture, confocal microscopy and PCR in routine hospital use for microbial keratitis diagnosis. Eye (2021). https://doi.org/10.1038/s41433-021-01812-7
  6. Lorenzo-Morales J, Khan NA, Walochnik J. An update on Acanthamoeba keratitis: diagnosis, pathogenesis and treatment. Parasite. 2015;22:10. doi: 10.1051/parasite/2015010. PMID: 25687209; PMCID: PMC4330640.

-Tim Kirtek is a fourth year AP/CP resident at UT Southwestern Medical Center in Dallas, Texas.

-Dominick Cavuoti is a professor at UT Southwestern Medical Center who practices Medical Microbiology, Cytology and Infectious Disease Pathology.

-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: A 75 Year Old Found Unresponsive

A 75 year old female with a past medical history of coronary artery disease, hypertension, pre-diabetes mellitus, chronic obstructive pulmonary disease, prior left lobe cavitary lesion of unknown etiology, and tobacco use presented to the ED after being found nonresponsive on the couch. Family reports the patient said she had emesis the night before and felt as if she had a “stomach bug”. MRI shows T2 hyperintensities in the right MCA distribution. CSF results as follows.

White Blood Cells300
Red Blood Cells12
Protein990
Glucose79
Cryptococcal antigenNegative
Fungal cultureNo fungi isolated
HSVNegative

Laboratory findings

CSF was sent to the microbiology lab for bacterial and fungal smears and cultures. No fungi were identified. Cryptococcal antigen was negative. HSV was also negative. CSF Gram stain shows gram positive bacilli. CSF culture showed a small, white, smooth, translucent appearance on sheep blood agar. In semi-solid agar after overnight incubation at room temperature, an umbrella shaped pattern of motility was seen. The organism was identified as Listeria monocytogenes by MALDI-TOF mass spectrometry.

Image 1. Listeria monocytogenes on sheep blood agar.
Image 2. Listeria monocytogenes showing “umbrella zone” pattern of motility on semi-solid agar.

Discussion

Listeria spp. is a genus of gram positive, aerobic, facultative intracellular, catalase positive bacteria. Listeria monocytogenes is a common colonizer in the environment (animals, soil, vegetable matter) and occasionally colonizes the human gastrointestinal tract. Listeria prefers colder environments and can be found as a food contaminant, most notably in milk, raw vegetables, cheese, and meats. In addition, colonized mothers can pass Listeria monocytogenes to the fetus.1

Listeria monocytogenes has 3 notable virulence factors:2

  1. Listeriololysin O: a hemolytic toxin that allows for survival within phagocytes
  2. Act A: induces actin polymerization that facilitate cell-to-cell spread
  3. Siderophores: organisms capable of scavenging iron from human transferrin to enhance cell growth

Neonates, immunocompromised individuals, and the elderly are more likely to acquire infection. Infection can present as bacteremia and CNS infections including meningitis, encephalitis, brain abscesses, and spinal cord infections. Listeria monocytogenes is the 3rd most common cause of meningitis behind Streptococcus pneumoniae and Neisseria Meningitidis. In neonates, an in-utero infection can cause granulomatous infantisepticum leading to systemic infection and stillbirth.3 Listeria monocytogenes can also present as gastroenteritis.

References

  1. Allerberger F. Listeria: growth, phenotypic differentiation and molecular microbiology. FEMS Immunol Med Microbiol. 2003;35(3):183-189. doi:10.1016/S0928-8244(02)00447-9
  2. Bailey & Scott’s Diagnostic Microbiology – Elsevier eBook on VitalSource, 14th Edition – 9780323433792. https://evolve.elsevier.com/cs/product/9780323433792?role=student
  3. Engelen-Lee JY, Koopmans MM, Brouwer MC, Aronica E, van de Beek D. Histopathology of Listeria Meningitis. Journal of Neuropathology & Experimental Neurology. 2018;77(10):950-957. doi:10.1093/jnen/nly077

-Nicholas Taylor, DO 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: A 35 Year Old Male with Left Leg Cellulitis

Clinical History

A 35 year old male with chronic bilateral lower extremity lymphedema due to obesity presented with a one-week history of subjective fevers and malaise with associated left lower extremity pain, swelling and erythema. The left leg was markedly edematous with erythema present above the knee down. The leg was tender to palpation, and multiple ruptured bullae and areas of severe desquamation with excessive serous drainage were observed. Importantly, no areas of purulence were noted (Image 2). A clinical diagnosis of severe non-purulent cellulitis was made, and the patient was admitted for parenteral antibiotic therapy of vancomycin and piperacillin-tazobactam. Necrotizing fasciitis was ruled out based on imaging, and significant clinical improvement was seen after 5 days of intravenous antibiotics. The patient was transitioned to oral therapy with amoxicillin-clavulanic acid and doxycycline for a total of 14 days of antibiotics.

Laboratory Workup

During the admission, urinalysis revealed turbid urine with elevated protein (30 mg/dL), and 2+ blood with 5 RBC/HPF on microscopic examination. Given the presence of protein with microscopic hematuria, causes of glomerulonephritis were investigated. Workup revealed a markedly elevated anti-streptolysin O (ASO) titer of 5310 (0-330) and a total complement (CH50) level of 14, which was low given his age. Urine sediment examination revealed red blood cell casts (Image 3). These clinical and laboratory findings were consistent with post-streptococcal glomerulonephritis (PSGN) due to Streptococcus pyogenes skin and soft tissue infection.

 
Image 1. Colony appearance and biochemical testing of S. pyogenes. A) Typical gram positive cocci in chains characteristic of streptococci. B) Growth on Sheep’s Blood Agar of small, translucent colonies with a wide zone of beta-hemolysis indicative of S. pyogenes. C) Catalase-negative S. pyogenes (left) compared to catalase-positive S. aureus (right). D) PYR-positive S. pyogenes (left) compared to PYR-negative S. aureus (right).
Image 2. Left lower extremity at presentation.
Image 3: Red blood cell cast seen in urine sediment.

Discussion

Streptococcus pyogenes are gram positive bacteria that appear in pairs and/or chains by microscopy (Image 1A). In culture, these organisms produce relatively small colonies which elaborate a large zone of beta hemolysis on blood agar plates; colonies are translucent with smooth edges (Image 1B). The beta-hemolytic activity of S. pyogenes is due to the activity of two hemolysins: Streptolysin-S (oxygen-stabile) and Streptolysin-O (oxygen-labile). S. pyogenes is the primary organism which expresses the Lancefield Group A carbohydrate antigen. Less frequently encountered strains of S. anginosus and S. dysgalactiae subsp. equisimilis may also express this antigen, so biochemical identification of S. pyogenes may be helpful for a definitive diagnosis. MALDI-TOF MS may also fail to discriminate between S. pyogenes and closely related β-hemolytic streptococci (including S. dysgalactiae and S. canis), necessitating adjunctive biochemical testing. Like other streptococci, S. pyogenes is catalase negative (Image 1C). Unlike other beta-hemolytic streptococci, S. pyogenes expresses pyrrolidonyl arylamidase (PYR) making this test a rapid and useful adjunctive diagnostic tool (Figure 1D). Bacitracin susceptibility was used historically but has been largely replaced by PYR testing due to concerns over specificity and prolonged turnaround time.

Globally, S. pyogenes is responsible for a large percentage of infection-related morbidity and mortality. The organism colonizes the skin and the nasopharynx of humans, but most colonized individuals do not develop active disease. Colonization however can lead to infection or dissemination to susceptible individuals. S. pyogenes infections exhibit a diverse range of clinical manifestations which can include pharyngitis, impetigo, erysipelas, cellulitis, necrotizing fasciitis, pyomyositis, streptococcal toxic shock syndrome, and bacteremia. S. pyogenes remains susceptible to penicillin, making β-lactams first-line drugs of choice for management. Conversely, rising levels of macrolide, lincomycin, tetracycline, and fluoroquinolone resistance has been observed. Susceptibility testing may be warranted if these agents are to be used, most often in the cases of severe penicillin allergy.

S. pyogenes infection can be complicated by multiple post-infectious immune-mediated sequelae including PSGN and rheumatic fever. Post-Streptococcus glomerulonephritis (PSGN) has a global incidence of > 470,000 individuals per year and occurs due to the deposition of immune complexes in the glomeruli resulting from previous S. pyogenes pharyngitis or soft tissue infection (as seen in this case). Typical clinical presentation of PSGN includes hematuria, proteinuria, edema, hypertension, elevated serum creatinine levels, hypocomplementemia, and general malaise. The elevated ASO titer (5310) was diagnostic of an S. pyogenes acute infection as the cause of this patient’s cellulitis. The development of proteinuria and hematuria following infection further supports a clinical diagnosis of PSGN. Treatment of PSGN is largely supportive with the focus on management of the underlying infection. Most individuals with kidney failure from PSGN recover to baseline renal function; however, there may be a link between PSGN and the later development of chronic kidney disease/end-stage renal disease.

References

  1. De la Maza LM, Pezzlo MT, Bittencourt CE, Peterson EM. 2020. Color Atlas of Medical Bacteriology, 3rd edition. ASM Press. Pg. 11-23
  2. Madaio MP, Harrington JT. 2001. The diagnosis of glomerular diseases: acute glomerulonephritis and the nephrotic syndrome. Arch Intern Med. 161(1):Pg. 25-34. doi: 10.1001/archinte.161.1.25.
  3. Stevens DL, Bisno AL, Chambers HF, Dellinger EP, Goldstein EJC, Gorbach SL, Hirschmann JV, Kaplan SL, Montoya JG, Wade JC. 2014. Practice Guidelines for the Diagnosis and Management of Skin and Soft Tissue Infections: 2014 Update by the Infectious Diseases Society of America. Clin Infect Dis. 59(2): Pg. e10-e52, https://doi.org/10.1093/cid/ciu296.
  4. Walker MJ, Barnett TC, McArthur JD, Cole JN, Gillen CM, Henningham A, Sriprakash KS, Sanderson-Smith ML, Nizet V. 2014. Disease manifestations and pathogenic mechanisms of Group A Streptococcus. Clin Microbiol Rev. (2): Pg. 264-301. doi: 10.1128/CMR.00101-13.
  5. Wong CH, Khin LW, Heng KS, Tan KC, Low CO. 2014. The LRINEC (Laboratory Risk Indicator for Necrotizing Fasciitis) score: a tool for distinguishing necrotizing fasciitis from other soft tissue infections. Crit Care Med. 32(7): Pg. 1535-41. doi: 10.1097/01.ccm.0000129486.35458.7d.

-John Markantonis, DO is the former Medical Microbiology fellow at UT Southwestern and has recently completed his clinical pathology residency. He is also interested in Transfusion Medicine and parasitic diseases.

-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: Genotypic-to-phenotypic Discordant Results

Case History

Scenario 1: A 51 year old male with a history of diabetes, hypertension, coronary artery disease, gastric ulcer, chronic kidney disease and bilateral below knee amputation presented with epigastric pain, nausea, and vomiting. He was febrile and tachycardic. Computerized scan showed ascending/ transverse colitis and cholelithiasis. Blood cultures grew gram negative rods; the Biofire BCIDv2 panel reported Enterobacter cloacae with no genotypic, resistance markers detected. Phenotypic antimicrobial susceptibility testing (AST) from the Microscan Walkaway revealed resistance to ertapenem (>1mg/ml) but susceptibility to meropenem (£ 1mg/ml). Additionally, the isolate was resistant to 3rd-generation cephalosporins, fluoroquinolones, and intermediate-resistant to tetracyclines. Identification was confirmed by the MALDI-TOF MS upon growth on agar plates. The isolate was subbed with a meropenem disk to select for carbapenem resistance for further confirmatory testing. A Cepheid Carba-R test was ran on a sweep of the isolate growing near the carbapenem disk, which resulted in no carbapenemases detected. Results from E-tests with meropenem and ertapenem were consistent with original phenotypic result. Here, we reported the discrepant phenotypic result and genotypic results as is.

Image 1. Phenotypic testing results (E-test) for meropenem (MP,left) and ertapenem (ETP, right) of Enterobacter cloacae isolate described in scenario 1. E-test results were consistent with original phenotypic results which also identified the isolate as meropenem susceptible and ertapenem resistant. (Photo credit: Gizachew Demessie, Lead Tech, George Washington Hospital.)

Scenario 2: An 80 year old female underwent a Whipple procedure for a pancreatic mass. A wound culture was submitted from the operating room which grew both Streptococcus anginosus and Enterobacter cloacae complex. Phenotypic AST for the E. cloacae revealed susceptibility to ertapenem (≤0.5 mg/ml) but resistance to meropenem (4 mg/ml). The isolate was pan-susceptible to other drug classes (aside from intrinsic resistance). Similar to Case 1 above, identification was confirmed by the MALDI-TOF MS and the isolate was subcultured with selective pressure. A Cepheid Carba-R test did not detect any carbapenemases. However, upon repeating a phenotypic test, both ertapenem and meropenem were susceptible. Our investigation here led to the avoidance of reporting an incorrect phenotypic AST result.

Discussion

Genotype-to-phenotype discrepancies may occur in antimicrobial susceptibility testing. For example, an antimicrobial resistance (AMR) gene may be detected in a phenotypically susceptible isolate or an AMR gene may not be detected in a phenotypically resistant isolate. Such discordant results should be investigated so appropriate antimicrobial therapy is used on these patients. This leads us to an important question “What can laboratories do to solve these discrepancies?”

The first step in detection of discrepancies requires educating and teaching the lab staff to be vigilant in looking for odd susceptibility patterns (from results within a drug class and also the overall AST profile). Next, check if there was pure isolation of the organism on the purity plate; if not, each individual isolate should be subbed, identified and re-tested on both genotypic and phenotypic platforms. Of note, subbing the bacteria under selective antibiotic pressure (e.g. growing the isolate on agar plate with an antibiotic disk) can increase the potential of detecting resistance. Alternative methods (e.g. CarbaNP, mCIM, etc) could be considered if one is looking into specific resistant mechanisms. Due diligence in checking for clerical, transcription errors and contamination on equipment, especially when there is a consistent pattern of detection for a specific molecular target, is highly recommended. As such, a lab should maintain constant communication with the test manufacturer in case there are issues with batches or lots of reagents.1,2

While these rapid, genotypic panels tend to include the more common AMR mechanisms, there are still other mechanisms of resistance not on the panels. For gram negatives, AMR mechanisms such as AmpC beta-lactamases, porin mutations, efflux pumps and rare carbapenemases such as GES, IMI, and SME types are typically not included.3 Additionally, although the gene blaCTX-M is used as a marker for Extended Spectrum Beta-Lactamases (ESBL), different variants of ESBLs confer different cephalosporin (e.g. 3rd and 4th generation) phenotypes.4 A heteroresistant subpopulation, decreased or lack of expression of an AMR gene may also be potential explanations.

If a discrepancy is not resolved, it is suggested to report the isolate as resistant. If both the discrepant genotypic and phenotypic results are reported, one should consider recommending an infectious diseases consult or to contact the antimicrobial stewardship team.1 Additional information and suggested laboratory workflow can be found in Appendix H of the M100 guidelines from the Clinical Laboratory and Standards Institute.2 While molecular AMR approaches have many advantages such as a shorter turnaround time, phenotypic susceptibility testing can still offer valuable clinical information.5

  1. CLSI. Performance Standards for Antimicrobial Susceptibility Test. CLSI supplement M100. Wayne, PA: Clinical and Laboratory Standards Institute; 2022, Edition 32
  2. Yee R, Dien Bard J, Simner PJ. The Genotype-to-Phenotype Dilemma: How Should Laboratories Approach Discordant Susceptibility Results? J Clin Microbiol. 2021 May 19;59(6):e00138-20.
  3. Tamma PD, Sharara SL, Pana ZD, Amoah J, Fisher SL, Tekle T, Doi Y, Simner PJ. 2019. Molecular epidemiology of ceftriaxone non-susceptible Enterobacterales isolates in an academic medical center in the United States. Open Forum Infect Dis 6:ofz353.
  4. Paterson DL, Bonomo RA. 2005. Extended-spectrum beta-lactamases: a clinical update. Clin Microbiol Rev 18:657–686.
  5. Dien Bard J, Lee F. 2018. Why can’t we just use PCR? The role of genotypic versus phenotypic testing for antimicrobial resistance testing. Clin Microbiol Newsl 40:87–95. 10.1016/j.clinmicnews.2018.05.003. 

Rami Abdulbaki, MD is a Pathology Resident (PGY-3) at The George Washington University Hospital. His academic interest includes hematopathology and molecular 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: How to “Pin” a Diagnosis

Case History

A 7 year old female presented to the emergency department with left sided abdominal pain and a temperature of 103 degrees Fahrenheit. Labs drawn showed mild leukocytosis with a CT scan suggestive of acute appendicitis. The patient underwent uncomplicated appendectomy with no complication. Gross examination of the appendix revealed an unremarkable, non-perforated serosa and a fecalith within the lumen. Representative tissue sections submitted for microscopic analysis per grossing policy. The findings below led to the submission of the entire appendix to be evaluated.

Figure 1. Low power image of an appendix demonstrating mild acute inflammation, lymphoid hyperplasia and congestion.

Figure 2. High power image, Cross-section of an adult female E. vermicularis from the same specimen shown in Figure 1. Adherent to the appendiceal surface. Note the presence of the alae (blue arrow), and the presence of almond shaped eggs (red arrow).

Discussion

The nematode Enterobius vermicularis, widely known as the human pinworm, is one of the most common parasitic worm infections today in the United States, infecting approximately 40 million people. The patient population is often children who are infected via fecal-oral transmission, with autoinfection being common. Humans are the only known host of this nematode. Once E. vermicularis embryonated oocytes are ingested, the larvae hatch and inhabit the gastrointestinal system. At night, the larvae migrate down to the anus, lay their eggs, and the cycle recurs.

The clinical presentation can be asymptomatic or can present with perianal pruritus at night, which can be explained via the life cycle of the parasite as stated above. The method of choice for diagnosing E. Vermicularis is microscopic examination of the eggs via cellulose tape slide test. A piece of scotch tape collects the eggs near the perianal area of the patient, which is then used for analysis and identification of the eggs. Microscopically, E. Vermicularis can be identified by the spines or ‘alas’ on the surface with oval shaped, thick capsuled oocytes within, seen in figure 2. To improve the sensitivity of the scotch tape test, it is best to do this test in the early morning, when there is an increased chance of sampling the eggs.

Rarely, is this worm associated with any severe symptoms but patients can present with abdominal pain, suggesting intestinal obstruction, extra intestinal manifestations like vulvovaginitis, or appendicitis. The relationship between E. Vermicularis and appendicitis is up for debate as to whether there is a causative relationship or if it is an incidental finding seen within appendicitis. Regardless of the relationship, once a diagnosis of Enterobius vermicularis is made, treatment with an anthelmintic needs to be given to the patient, such as Albendazole or Pyrantel Pamoate. In addition, treatment for everyone in the household needs to be considered in confirmed cases of infection.

Routine surgical specimens, such as appendices, can perhaps be overlooked once acute inflammation is noted. It is important to be able to identify organisms, such as pinworms, on such specimens to get the patient the appropriate treatment.

References

  1. https://www.cdc.gov/dpdx/enterobiasis/index.html.
  2. https://www.sciencedirect.com/science/article/pii/S204908012030412X
  3. https://www.uptodate.com/contents/enterobiasis-pinworm-and-trichuriasis-whipworm?search=enterobius%20vermicularis&source=search_result&selectedTitle=1~32&usage_type=default&display_rank=1#H12

-Alexandra Medeiros, MD, is a first year anatomic and clinical pathology resident at Medical College of Georgia at Augusta University. Her academic interests include Forensic pathology, and surgical pathology.

-Hasan Samra, MD, is the Director of Clinical Microbiology at Augusta University and an Assistant Professor at the Medical College of Georgia.

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.