Microbiology Case Study: Not An Ordinary Sore Throat, but One Accompanied by Headache 

Case History

An 18 year old healthy female presented to the emergency department of a tertiary care hospital in Minnesota for headache, vomiting, and sore throat. She did not have any significant past medical history. Due to meningitis concerns, lumbar puncture and head computed tomography (CT) imaging were performed. The CT scan showed an accumulation of fluid in the posterior right frontal sinus with scattered mucosal thickening. However, her cerebrospinal fluid (CSF) profile was insignificant, with normal protein and glucose levels. CSF culture was ordered, and two sets of blood cultures were drawn. 

Based on the examination and presenting symptoms, pharyngitis was suspected, and she was discharged with Amoxicillin (500mg Q6H for five days). However, her strep throat screening returned negative. Her blood culture was negative. CSF culture was also negative. Cryptococcal antigen and Enterovirus PCR were performed; however, both results were negative.

She returned to the ED two days later for a worsening headache and newly developed photophobia. Additional history revealed that she went swimming in a lake two weeks prior to her first presentation at the ED. Her CSF was sent for the Ova and Parasite (O&P) exam for suspicious parasitic meningitis. The CSF O&P test was negative. CSF PCR for amoeba was also performed at a reference laboratory, and the results came back positive with Balamuthia mandrillaris. The patient was then given flucytosine, fluconazole, and azithromycin. 

Figure 1. Photo Credit CDC: Balamuthia – free living parasite observed under light microscopy.


Balamuthia mandrillaris belongs to a group of free-living amoebae, including Acanthamoeba species and Naegleriafowleri, that cause fatal encephalitis.1 Balamuthia mandrillaris is the only known species of the genus Balamuthia that causes infections in humans. Encephalitis caused by B. mandrillaris is known as granulomatous amoebic encephalitis (GAE). GAE is characterized as a subacute to a chronic infection that can last several months to years.2 GAE differs from primary amoebic meningoencephalitis (PAM) caused by Naegleria fowleri, which typically causes an acute onset lasting a few days. 

While ecological niches of B. mandrillaris are not well understood, they have been reported to be isolated from dust, soil, and water.r Both trophozoite and cyst forms can enter the body through the nasal passage or ulcerated/broken skin; however, the trophozoite stage causes associated disease manifestation and represents a diagnostic stage.2 

Brain-eating amoebas are traditionally difficult to diagnose. Hematology and chemistry profiles of CSF of affected individuals are generally unremarkable, although, sometimes, increased monocytes and lymphocytes, along with increased protein levels, are seen in some cases of GAE.3 

The most common method of laboratory diagnosis of B. mandrillaris is a microscopic examination of CSF wet mount (Figure 1) or via immunohistochemical staining of CSF or brain biopsy.1 With advancements in technology, species-specific nucleic acid amplification tests (NAAT) can be performed to diagnose B. mandrillaris infection accurately. However, there is no commercially available NAAT for the free-living amoeba. Only very few laboratories, such as State departments of health laboratories, Centers for Disease Control (CDC) and Prevention, or commercial reference laboratories, develop these tests as a laboratory-developed test (LDT). Histological assessment of biopsies from brain lesions may reveal tumor-like appearance or perivascular monocytic necrosis of affected areas.1 While there have been significant technological advancements, the prognosis stays at less than a 5% survival rate,6 with only roughly 25% of cases diagnosed antemortem. One possible reason for delayed laboratory diagnosis is the challenges in performing the microscopic examination in clinical microbiology laboratories since it requires expertise for accurate identification of the organism. Additionally, most clinical microbiology laboratories do not readily have an in-house LDT for free-living amoeba NAAT. Therefore, the turnaround time for diagnosing B. mandrillaris or any free-living amoeba is typically longer when specimens have to be sent out to reference laboratories. 

Diagnosis of B. mandrillaris encephalitis solely based on clinical symptoms is often challenging due to similar presentation in other causes of infectious encephalitis. B. mandrillaris can affect both immunocompetent and immunocompromised individuals.1,3,4 The first B. mandrillaris case was reported in a deceased baboon in the San Diego Zoo in 1986.1  The majority of patients were diagnosed postmortem.1 While most B. mandrillaris infections are actively acquired through nasal passages or skin penetration, rare post-mortem cases of passive transfer of the organism from organ transplantation have been reported.6 With technological advancement, there have been successes in pre-mortem diagnoses in recent years.1,4 According to the known cases, individuals of Latin American origin are more likely to contract the disease; it is unknown if it is due to increased exposure or a genetic predisposition.1  Similar to other free-living amoebae, B. mandrillaris can be generally found in warmer climates or tropical regions. Of approximately two hundred cases reported worldwide, about 34 were reported in Latin America, from Mexico to Brazil, while some were from Japan, New Zealand, England, and other European countries. The Southwestern United States also contributes 30 cases, mostly in Arizona, Texas, and California.1  In the United States, there have only been 109 cases directly reported to CDC from 1974 to 2016.2,7  We believe that this is the first case of Balamuthia reported in Minnesota. The number of exact cases would be difficult to be determined due to misdiagnosis and rare occurrence of the disease or cases not reported to CDC or the state department of health. 

While investigational drugs for B. mandrillaris GAE are in development, combination therapy of flucytosine, fluconazole, pentamidine, and azithromycin or clarithromycin has shown successes.2 Our patient was successfully treated with flucytosine, fluconazole, and azithromycin. 


  1. Matin A, Siddiqui R, Jayasekera S, Khan NA. Increasing importance of Balamuthia mandrillaris. Clin Microbiol Rev. 2008 Jul;21(3):435-48. doi: 10.1128/CMR.00056-07. PMID: 18625680; PMCID: PMC2493082. 
  2. Centers for Disease Control and Prevention. (2019, August 23). CDC – Dpdx – free Living Amebic Infections. Centers for Disease Control and Prevention. https://www.cdc.gov/dpdx/freelivingamebic/index.html.
  3. Kofman A, Guarner J. Free Living Amoebic Infections: Review. J Clin Microbiol. 2021 Jun 16:JCM0022821. doi: 10.1128/JCM.00228-21. Epub ahead of print. PMID: 34133896.
  4. Pietrucha-Dilanchian, P., Chan, J. C., Castellano-Sanchez, A., Hirzel, A., Laowansiri, P., Tuda, C., Visvesvara, G. S., Qvarnstrom, Y., & Ratzan, K. R. (2011). Balamuthia mandrillaris And Acanthamoeba Amebic Encephalitis With Neurotoxoplasmosis Coinfection in a patient with Advanced HIV Infection. Journal of Clinical Microbiology, 50(3), 1128–1131.
  5. Ong TYY, Khan NA, Siddiqui R. 2017. Brain-eating amoebae: predilection sites in the brain and disease outcome. J Clin Microbiol 55:1989 –1997. https://doi.org/10.1128/JCM. 02300-16.
  6. Centers for Disease Control and Prevention. 2011. Balamuthia mandrillaris transmitted through organ transplantation—Mississippi, 2009. Am J Trans-plant 11:173–176. https://doi.org/10.1111/j.1600-6143.2010.03395_1.x.
  7. Jennifer R Cope, Janet Landa, Hannah Nethercut, Sarah A Collier, Carol Glaser, Melanie Moser, Raghuveer Puttagunta, Jonathan S Yoder, Ibne K Ali, Sharon L Roy, The Epidemiology and Clinical Features of Balamuthia mandrillaris Disease in the United States, 1974–2016, Clinical Infectious Diseases, Volume 68, Issue 11, 1 June 2019, Pages 1815–1822, https://doi.org/10.1093/cid/ciy813

-Alejandro Soto, MLS (ASCP)CM is a junior medical technologist who is passionate about clinical microbiology.

-Phyu M. Thwe, Ph.D., D(ABMM), MLS(ASCP)CM is Microbiology Technical Director at Allina Health Laboratory in Minneapolis, MN. She completed her CPEP microbiology fellowship at the University of Texas Medical Branch in Galveston, TX. Her interest includes appropriate test utilization and extra-pulmonary tuberculosis.

Microbiology Case Study: A 62 Year Old Male with Altered Mental Status

Case Description

A 62 year old male with unknown past medical history was dropped off at the emergency department by EMS after being found altered with concern for IV drug use. On presentation he was febrile to 104.5o F, tachycardic, and although he was initially responsive, his mental status deteriorated. Labs were drawn and broad-spectrum antibiotic coverage with vancomycin, cefepime, and metronidazole was initiated in the ED. He then had a tonic-clonic seizure event and was given intravenous levetiracetam. A CT brain showed a right inferior temporal lobe lesion, initially interpreted as likely glioblastoma multiforme, causing subfalcine and uncal herniation. MRI revealed a ring-enhancing mass measuring 3 cm x 3 cm x 3 cm in the right temporal lobe with significant surrounding edema. CT of the temporal bones also revealed right mastoiditis (Figures 1 and 2).

Figure 1. Coronal T1 post-contrast MRI demonstrating the ring-enhancing mass in the right temporal lobe (arrow).
Figure 2. CT of temporal bones with IV contrast demonstrating opacification of the right mastoid air cells and abnormal soft tissue within the epitympanum (arrow).

Neurosurgery evacuated 17 mL of fluid from the mass and a ventricular drain was placed. Gram stain of the evacuated fluid identified many white blood cells and few gram-positive cocci in pairs, chains and clusters (Figure 3). Postoperatively, the patient was mechanically ventilated and medicines were used to support his blood pressure in the ICU. Broad-spectrum antibiotics were continued for CNS penetration and activity against possible oral flora.

Figure 3. Representative Gram stain of the pus drained from the abscess. Multiple couplets of lancet-shaped gram positive organisms identified. Slight halo around the bacteria suggests the presence of a capsule.

An aerobic culture of drained contents from the brain ultimately grew characteristic alpha-hemolytic colonies with central umbilication (Figure 4) which were subsequently identified by MALDI-TOF and optochin disk as Streptococcus pneumoniae. Admission blood cultures also grew Streptococcus pneumoniae with characteristic “bullet-” or “lancet-shaped” gram-positive cocci in pairs on Gram stain. Fungal and acid-fast bacillus cultures had no growth. Following susceptibility testing; antibiotic coverage was narrowed to IV Penicillin G.

Figure 4: Representative, archival image of a blood agar plate with alpha-hemolytic, centrally umbilicated colonies and optochin susceptibility (P disk).

The patient remained unresponsive and required continued intensive medical support. Although blood cultures were sterilized, he continued to have fevers and persistent leukocytosis. Gram stain of ventricular drainage re-demonstrated gram positive diplococci. The patient was transitioned to comfort care and expired on day 5 of hospitalization, the cause of death was sepsis.


This case of a brain abscess demonstrates an unusual intracranial complication of Streptococcus pneumoniae. S. pneumoniae (or pneumococcus) is a commensal of the upper respiratory tract (URT) and important opportunistic pathogen. Up to 65% of children and less than 10% of adults are colonized by S. pneumoniae. Dissemination of S. pneumoniae beyond its niche in the nasal mucosa leads to a spectrum of disease including lobar pneumonia, meningitis, sepsis, sinus infections and middle ear infections.1 Local dissemination of S. pneumoniae to the central nervous system (CNS) is the most common intracranial complication of otitis media and mastoiditis. These patients can present with fulminant “otogenic” meningitis. About a third of these cases require myringotomy or mastoidectomy.2 Focal parenchymal brain infection by pneumococcus, however, is uncommon.

This patient presented with signs of mass effect due to a large temporal lobe abscess warranting emergent neurosurgery. Broadly, focal parenchymal brain infections arise either by hematogenous dissemination of organisms or contiguous spread from an adjacent infection. The age, immune status, and any underlying disease present in the patient help predict the pathogen. Brain imaging is also helpful. Hematogenous spread, usually from endocarditis, tends to produce multiple lesions at the grey-white matter junction,3 while direct seeding causes solitary lesions.4,5 In this older patient with a relatively intact immune system and a possible history of intravenous drug use, hematogenous spread of bacteria was considered. However, a large single lesion in the temporal lobe with a plausible adjacent nidus (opacified mastoid air cells) is most consistent with contiguous spread.

A wide range of organisms should be considered when evaluating brain abscesses, though S. pneumoniae is a relatively rare culprit. A meta-analysis of 9,699 patients with brain abscesses found that S. pneumoniae was isolated from only 2.4%.6 The most common organisms were streptococci of the viridans group (34%) and Staphylococcus spp., most commonly S. aureus (18%). Even among patients with otogenic intracranial abscesses, S. pneumoniae is rarely implicated. Interestingly, the pathogen most frequently isolated from otogenic brain abscesses is Proteus mirabilis. 7,8

Once S. pneumoniae was identified, susceptibility testing was required to rule out acquired resistance to beta lactam and cephalosporin antibiotics, which is mediated by altered penicillin-binding proteins (PBPs).9 A more stringent susceptibility minimal inhibitory concentration (MIC) breakpoint applies to S. pneumoniae meningitis than other infections to account for drug distribution into the CNS.10 The hospital antibiogram reports that 97% of S. pneumoniae isolates are susceptible to Penicillin at MICs acceptable for treating non-meningitis infection but only 53% are susceptible at MICs for meningitis. Furthermore, 3.3% of all strains reported in the United States between 2001 and 2005 were also significantly resistant to ceftriaxone.11 This patient was covered by broad spectrum antibiotics until susceptibility testing demonstrated sensitivity to both penicillin and ceftriaxone.


1          Weiser, J. N., Ferreira, D. M. & Paton, J. C. Streptococcus pneumoniae: transmission, colonization and invasion. Nat Rev Microbiol 16, 355-367, doi:10.1038/s41579-018-0001-8 (2018).

2          Kaplan, D. M., Gluck, O., Kraus, M., Slovik, Y. & Juwad, H. Acute bacterial meningitis caused by acute otitis media in adults: A series of 12 patients. Ear Nose Throat J 96, 20-28 (2017).

3          Bakshi, R. et al. Cranial magnetic resonance imaging findings in bacterial endocarditis: the neuroimaging spectrum of septic brain embolization demonstrated in twelve patients. J Neuroimaging 9, 78-84, doi:10.1111/jon19999278 (1999).

4          Brouwer, M. C., Tunkel, A. R., McKhann, G. M., 2nd & van de Beek, D. Brain abscess. N Engl J Med 371, 447-456, doi:10.1056/NEJMra1301635 (2014).

5          Miller, J. M. et al. A Guide to Utilization of the Microbiology Laboratory for Diagnosis of Infectious Diseases: 2018 Update by the Infectious Diseases Society of America and the American Society for Microbiology. Clin Infect Dis 67, e1-e94, doi:10.1093/cid/ciy381 (2018).

6          Brouwer, M. C., Coutinho, J. M. & van de Beek, D. Clinical characteristics and outcome of brain abscess: systematic review and meta-analysis. Neurology 82, 806-813, doi:10.1212/WNL.0000000000000172 (2014).

7          Duarte, M. J. et al. Otogenic brain abscesses: A systematic review. Laryngoscope Investig Otolaryngol 3, 198-208, doi:10.1002/lio2.150 (2018).

8          Kangsanarak, J., Fooanant, S., Ruckphaopunt, K., Navacharoen, N. & Teotrakul, S. Extracranial and intracranial complications of suppurative otitis media. Report of 102 cases. J Laryngol Otol 107, 999-1004, doi:10.1017/s0022215100125095 (1993).

9          Chen, L. F., Chopra, T. & Kaye, K. S. Pathogens resistant to antibacterial agents. Infect Dis Clin North Am 23, 817-845, vii, doi:10.1016/j.idc.2009.06.002 (2009).

10        Weinstein, M. P., Klugman, K. P. & Jones, R. N. Rationale for revised penicillin susceptibility breakpoints versus Streptococcus pneumoniae: coping with antimicrobial susceptibility in an era of resistance. Clin Infect Dis 48, 1596-1600, doi:10.1086/598975 (2009).

11        Sahm, D. F. et al. Tracking resistance among bacterial respiratory tract pathogens: summary of findings of the TRUST Surveillance Initiative, 2001-2005. Postgrad Med 120, 8-15, doi:10.3810/pgm.2008.09.suppl52.279 (2008).

Miles Black, Ph.D. is a fourth-year medical student in the Medical Scientist Training Program at UT Southwestern Medical Center. His background is in enzyme biochemistry and Legionella pathogenesis.

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.

Dominick Cavuoti, D.O. is a professor of Pathology at UT Southwestern Medical Center who specializes in Medical Microbiology, ID Pathology and Cytology.

-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.

Pre-Analytic factors in NGS: Recognizing Contamination

Pre-analytic factors contribute to over 70% of laboratory errors. The most common examples include missing test request, wrong/ missing ID, contamination from collection site, hemolyzed/ clotted/ insufficient sample, wrong container or wrong transport conditions. 

In our NGS lab, pre-analytic factors rarely arise. Sometimes an incorrect slide was sent, so then the tumor and germline DNA don’t match. We do perform many rounds of PCR (total 30-35 cycles), so contamination of amplified PCR products could be a concern. Usually, our volume is low enough that contamination isn’t a significant issue and not one we have noticed.

Multiple mutations- A COVID-19 Hybrid?

In a batch of COVID-19 whole genome sequencing samples from early July, I noticed some interesting mutations. Mutations from the Alpha and Delta variants were showing up in the same specimen. At first I thought this could mean a hybrid virus. Another possibility was co-infection in the same patient. And lastly, the specimen could just be contaminated. The best way to resolve this issue is to check the “phase” of the viral variant sequencing reads. The phase refers to the single pieces of DNA that are read by the sequencer. They are normally short (75-150bp in length), but if you can see whether the variants occur on the same read or only on different reads it rules out the possibility of a hybrid virus.

Image 1. Hybrid (or chimeric) cat.

Determining the difference between a hybrid or 2 viral genomes

The difficulty is finding a location within both the Alpha and Delta variants that have mutations close to each other. One region exists near a deletion at amino acid 144 (Alpha) and amino acids 157-158 (Delta).

Figure 1. Characteristic mutations of Alpha (top) and Delta (bottom) variants. The deletions (triangle sign) are located close to each other, so were evaluated for phase.
Figure 2. Integrated Genome View (IGV) display of mutations. 6 base pair deletion (bars next to 6) is characteristic of Delta while a 3 base pair deletion (bars next to 3) is characteristic of Alpha.

Tracing the source of the issue

As there were no overlaping reads with the 3 b.p. and 6 b.p. deletion, we concluded that this finding arose from 2 distinct viral genomes. As to whether the person was infected with both Alpha and Delta, we looked to the rest of our 96 well plate. A characteristic muation in Alpha spike protein is N501Y (it confers increased binding to ACE2R and increased infectivity). This mutation was found in several specimens of the Delta lineage, but in this case there was a much lower frequency of this variant compared to total reads. Also, several of the cases had high CT values to start with. Many of the contaminated samples also were near a variant that mapped strongly to Alpha and had a lower Ct value (CT=25). Mapping the location of the specmens to the plate showed close proximity to the authentic Alpha variant.

Figure 3. Red squares: PCR=B.1.1.7; others are PCR=B.1.1.7, but yellow highlighting indicates N501Y detected in WGS.

Luckily, before all WGS testing, we perform a targeted PCR, which I’ve mentioned in previous blog posts. This showed that only one sample (well F3) was B.1.1.7, and the rest were Delta variants. Thus we concluded we had experienced a case of contamination.

Since this time, we’ve had issues with the negative control having borderline positive levels of sequencing data that maps to Delta (now 100% of all cases and with low CT values). This is apparently a problem at multiple labs, but one that we are trying to address by looking at several root causes.

Pre-analytic concerns: Contamination sources

COVID-19 viral genome sequencing has found issues of contamination in several circumstances. We attribute this to a few reasons:

  • High viral load of some specimens (especially Delta)
  • Thin plastic covers that pop off easily, especially when taken out of the freezer
  • And a large volume of specimens being processed.

For an example of the plastic cover issue, you can see this picture below where the cold temperature of -80C causes the plastic film adhesive to come off quickly as it is removed from cold storage. It makes popping sounds and could aerosolize viral particles to other wells in the plate.

Image 2. 96 deep well plate of extracted RNA with a thin plastic cover that comes off as the plates is thawed.

We now use aluminum PCR plate covers that do not come off with freeze/ thaw transitions. Furthermore, we use a multi-channel pipette that pieces the cover to withdraw individual samples without exposing them to other wells.

We have also implemented bioinformatic QC metric cut-offs to determine where a cut-off for negative specimens should be. We decided on an ambiguity score <0.5. The results were consistent with previous findings where lineages could be assigned at a CT >30, but sometimes they failed as CT values increased. Negative controls were assigned a CT of 40 and all fell below the 0.5 cut-off. This has been a useful metric to be sure we are providing high quality results.

Concluding remarks

Pre-analytic factors impact every part of testing and as COVID-19 sequencing has shown, even the NGS lab tests are not immune to these challenges.

The targeted PCR test helped flag/ resolve several of the issues as they arose.

COVID-19 sequencing is still for research/ epidemiologic purposes and demonstrates the importance of rigorous clinical validations to mitigate issues such as carry-over.

References Lippi G, Chance JJ, Church S, Dazzi P, Fontana R, Giavarina D, et al. Preanalytical quality improvement: from dream to reality. Clin Chem Lab Med. 2011;49:1113–26.

-Jeff SoRelle, MD is Assistant Professor of Pathology at the University of Texas Southwestern Medical Center in Dallas, TX working in the Next Generation Sequencing lab. His research interests include the genetics of allergy, COVID-19 variant sequencing, and lab medicine of transgender healthcare. Follow him on Twitter @Jeff_SoRelle.

Microbiology Case Study: Salads, Stool, and Special Staining Studies

Case History

A woman in her 40s presented to her primary care physician in summer 2020 with mild abdominal pain, diarrhea, nausea, and headache. She experienced loose bowel movements 3 – 4 times per day for the past 18 days. She denied bloody stools, travel, consumption of raw or undercooked meats or unpasteurized dairy, contact with animals, or recent antibiotic use. SARS-CoV-2 PCR was negative. A stool sample was collected and sent for an enteric panel PCR (Salmonella, Shigella, Campylobacter, and Shiga toxin), a bacterial stool culture (Aeromonas, Plesiomonas, and Vibrio), and an ova and parasite (O&P) exam with a request to perform a modified acid-fast stain. While the enteric panel and bacterial stool culture were negative, the following organism was observed on the modified acid-fast stain (Image 1). This organism measured approximately 9 µm, was variably modified acid-fast, and had a wrinkled-cellophane appearance. The organism was identified as a Cyclospora cayetanensis oocyst. The patient later shared that she had consumed a bagged salad mix that was implicated in the ongoing Cyclospora outbreak.

Image 1. Cyclospora cayetanensis.

Cyclospora cayetanensis

Cyclospora cayetanensis, a coccidian protozoan, is transmitted through ingestion of food or water contaminated with infectious oocysts. While infected humans shed oocysts in their stool, these oocysts are unsporulated and non-infectious at the time of excretion. In order to sporulate and become infectious, these excreted oocysts must incubate in the environment for 7 – 15 days post-excretion. Due to the required incubation post-excretion, direct fecal-oral transmission cannot occur.

Endemic areas include Central and South America, Middle East, South East Asia, and the Indian subcontinent. In non-endemic areas, travelers make up a large proportion of cases. Local outbreaks in non-endemic areas are often due to contaminated food sources. Most commonly the source of these outbreaks arise from consumption of raw fruits and vegetables that are difficult to thoroughly clean. These include leafy green vegetables (salad mixes, lettuce), herbs (basil, cilantro), and raspberries. Moreover, Cyclospora is resistant to many disinfectants used in the food industry. As exposure to this parasite is through contaminated food and water, infected patients are also at risk for other food and waterborne parasites including Cryptosporidium.

Once infectious oocysts are ingested, symptoms are typically observed after a one week incubation. Clinical presentation of Cyclospora infection includes diarrhea, nausea, fatigue, low grade fever, and weight loss. Although Cyclospora causes infections in both immunocompromised and immunocompetent individuals, symptoms may be severe and prolonged in immunocompromised patients, particularly in HIV and AIDS patients. Children and elderly individuals are also at higher risk for severe disease. Trimethoprim-sulfamethoxazole is the standard treatment. If untreated, symptoms can last for 10 – 12 weeks and may exhibit a relapsing pattern.

Stool samples should be submitted to the clinical microbiology laboratory for microscopic and/or molecular studies. To increase recovery of the organism during intermittent or low burden shedding, multiple stool specimens should be submitted over 2 -3 days. When viewed under a UV fluorescent microscope, Cyclospora oocysts autofluoresce and appear blue or green. While safranin-based stains or UV fluorescent microscopic examination can be used, modified acid-fast staining is commonly performed for the microscopic identification of Cyclospora. Cyclospora oocysts are modified acid-fast variable and measure 8 – 10 µm in diameter, unlike Cryptosporidium oocysts which are modified acid-fast positive and measure 4 – 6 µm in diameter. It is important to alert the clinical microbiology lab if suspecting Cyclospora as stains used in routine O&P exam, including trichrome stains, are not effective in highlighting Cyclospora. Although lab developed tests and FDA cleared multiplex gastrointestinal pathogen panels including Cyclospora are available, molecular assays are not yet routinely used for identification of Cyclospora due limited widespread availability.


  1. Almeria S, Cinar HN, Dubey JP. Cyclospora cayetanensis and Cyclosporiasis: An Update. Microorganisms. 2019;7(9):317. Published 2019 Sep 4. doi:10.3390/microorganisms7090317
  2. Hadjilouka A, Tsaltas D. Cyclospora Cayetanensis-Major Outbreaks from Ready to Eat Fresh Fruits and Vegetables. Foods. 2020;9(11):1703. Published 2020 Nov 20. doi:10.3390/foods9111703
  3. Ortega YR, Sanchez R. Update on Cyclospora cayetanensis, a food-borne and waterborne parasite. Clin Microbiol Rev. 2010;23(1):218-234. doi:10.1128/CMR.00026-09
  4. Garcia LS. Diagnostic Medical Parasitology. 6th Edition. 2016.
  5. Casillas SM, Bennett C, Straily A. Notes from the Field: Multiple Cyclosporiasis Outbreaks — United States, 2018. MMWR Morb Mortal Wkly Rep 2018;67:1101–1102. DOI: http://dx.doi.org/10.15585/mmwr.mm6739a6

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.

The More Things Change: 3 Ways to Determine Lab Safety Culture

On which side of the aisle do you stand regarding the subject of change? Things change, or things never change? The only constant is change, or it’s always the same old thing? When it comes to the laboratory safety culture, there are some generally-accepted thoughts; change is difficult, change is slow, and change takes persistence and patience.

I’ve heard other things too- people hate change, or people like change as long as they get to be in charge of it. I do believe most of us like change. After all, we change our clothes, we re-arrange our furniture, we remodel a room in our home. It can be exciting, but the tables seem to turn if it’s a change that is forced upon us or that was not our decision (Think about all of the changes the pandemic forced upon us last year!). Changing your lab safety culture for the better can be difficult, but it can be done. First, however, you need to know the current culture and goings-on in your lab in order to be able to make a difference.

There are specific ways to determine the safety culture in your lab. An experienced safety professional can do it fairly quickly. For others, especially those who serve in multiple capacities (you know who you are- you’re in charge of lab safety but you’re also the lab manager, or the quality coordinator, or the POCT coordinator) – for you assessing the culture can be difficult, even with years of experience- because you have so many other things on your plate. That can hinder your ability to make quick assessments, but it will not hinder you completely from being able to make a true safety assessment.

To make an assessment you need to use three specific tools that you likely have at your disposal. These tools may come in a variety of forms.

Those who have followed my work for some time know about the tool “Safety Eyes.” This is a safety assessment tool I believe to be a “superpower” that we all have and need to develop. It is so powerful, in fact, that a developed user can make a fairly good and accurate safety assessment with a quick glance into the department. Performing a lab safety audit is also a very valuable tool that can give you much information about the department’s culture. Perform a complete audit at least annually, and follow-up on the results. Otherwise, you have wasted your time and resources.

A second important safety culture gauge is the use of a written or electronic safety culture assessment. You may be able to tell what’s going on visually and physically by the evidence of your eyes and safety audits, but this tool is a way to actually get into the heads of your staff. What do they think of the culture? What is their opinion of it? What do they think needs improvement, and how would they suggest making those changes? A safety culture assessment can be given to everyone, or it can be used for specific lab groups. Survey the lab staff, survey those responsible for safety, or survey lab leadership. You should perform a lab safety culture assessment at least annually, but it can be done more often as needed.

Lastly, you can use laboratory data that you already collect to see the current state of safety in the department. Analyzing the data you collect about the injuries, accidents and exposures in your laboratory can be very eye-opening, and if you share the data as safety education, you may be able to lower the number of these types of incidents. Look at the chemical and biological spills in the lab. Analyze how they happened and how to prevent a re-occurrence. If you’re the quality coordinator for your lab or system, you know about root cause and common cause analyses. The incidents that occur in the lab that generate a root cause investigation may not always be about lab safety, but it’s possible that investigations show safety is a key factor, and those results should be reviewed with the safety person in the lab.

There is much fact-gathering in the laboratory setting, even regarding the topic of safety. However, all of that data becomes worthless if there is no action taken with it. Audits, injury data, spill information – it can be very valuable information and it can all be used as tools to help you truly change your lab safety culture. If you use them properly, you can make a change, you can make a difference, and you might just end up on the correct side of the change aisle!

Dan Scungio, MT(ASCP), SLS, CQA (ASQ) has over 25 years experience as a certified medical technologist. Today he is the Laboratory Safety Officer for Sentara Healthcare, a system of seven hospitals and over 20 laboratories and draw sites in the Tidewater area of Virginia. He is also known as Dan the Lab Safety Man, a lab safety consultant, educator, and trainer.

Microbiology Case Study: A 60 Year Old Male with Dysuria

Case Description

A 60 year old Hispanic male with a past medical history significant for chronic pancreatitis, hypertension and cirrhosis was admitted with decompensated cirrhosis. He underwent paracentesis for ascites and subsequently developed a hematoma as a complication of the procedure which required embolization. During his 12-day long hospital stay, he also developed hypoxia due to volume overload that improved with diuresis. A Foley catheter was placed during his hospital stay which was removed prior to discharge. Weeks later, at a follow up appointment with urology, he complained of dysuria, very little urine during voiding and the sensation of incomplete bladder emptying. A clean catch urine culture was performed and grew >100,000 colonies of Escherichia coli. As shown in Table 1, the isolate was resistant to multiple classes of antibiotics including penicillins, cephalosporins, fluoroquinolones, one aminoglycoside (Tobramycin), Trimethoprim/Sulfamethoxazole, aztreonam and carbapenems (Ertapenem/Meropenem) making this isolate multi-drug resistant (MDR). Because of the resistance profile to the carbapenems, molecular testing for carbapenemase genes was performed and the New Delhi metallo-beta-lactamase (NDM-1) gene was detected. The patient was treated with nitrofurantoin for his symptomatic urinary tract infection (UTI).

Table 1. Antimicrobial susceptibility of this isolate of E. coli.


Escherichia coli is a gram negative, motile bacillus that is a normal constituent of the gastrointestinal tract and is one of the most common causes of uncomplicated UTI. Antimicrobial susceptibilities are nearly always performed because the isolates of E. coli can vary in resistance. E. coli do not have any intrinsic resistance to antibiotics other than penicillin; however, they can acquire resistance through numerous mechanisms including structural mutations and plasmid-borne genes that encode enzymes to various classes of antibiotics. One such plasmid-encoded enzyme is the NDM, which was identified in our patient’s isolate. Its resistance is the result of bacterial synthesis of a carbapenemase that deactivates carbapenems by breaking down the beta-lactam ring.1 In the United States, K. pneumoniae carbapenemase (KPC) is the most common, but other types carbapenemase enzymes have also been reported.1,2 NDM is uncommonly isolated in E. coli; it is more often identified in other gram negative bacteria including MDR Pseudomonas aeruginosa or Acinetobacter baumannii complex, which can cause, among other things, devastating nosocomial infections within a healthcare setting. Because these enzymes are on mobile elements, a patient can be colonized with one bacterial strain that carries the plasmid with the carbapenemase on it and transfer a copy of that plasmid to another bacterial strain, thereby conferring new carbapenem resistance to the new bacterium (e.g., P. aeruginosa with the NDM on a plasmid shares that plasmid with an E. coli). Carbapenem resistant Enterobacteriaceae (CRE) are of great importance in healthcare. Carbapenem resistance mediated by enzyme activity (e.g., KPC, NDM, OXA-48, etc), typically confers resistance to all beta lactams. Interestingly, NDM enzymes typically do not destroy aztreonam, a monobactam;3 however, it is common for bacteria to have multiple resistance genes, so NDM carrying strains can be resistant to aztreonam. Although these CRE isolates can cause significant morbidity and mortality when found in clinical samples including sputum or blood, luckily for our patient, he had an uncomplicated UTI and nitrofurantoin was susceptible.

-Limin Yang is a PGY-1 resident in Anatomic and Clinical Pathology at University of Texas Southwestern. She has varied interests including anatomic pathology specialties.

-Dominick Cavuoti is a professor of Anatomic and Clinical Pathology at UT Southwestern and active faculty on both Microbiology and Cytology services.

-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.

How to Rock a Risk Assessment

As a lab safety professional, we know performing risk assessments is an integral piece of managing a safety program. In fact, assessing risks and identifying hazards are considered the beginning steps that must be completed when approaching the management of any safety-related area. Risk assessments are the starting point for handling a bloodborne pathogens program, chemical hygiene, personal protective equipment, and many other lab matters. But how can you be sure they have been performed correctly, and how often should they be performed?

OSHA gives simple guidance on basic assessment of risk in the workplace. “The employer shall assess the workplace to determine if hazards are present, or are likely to be present, which necessitate the use of personal protective equipment (PPE).” They further state, “the employer shall verify that the required workplace hazard assessment has been performed through a written certification that identifies the workplace evaluated; the person certifying that the evaluation has been performed; the dates(s) of the hazard assessment; and which identifies the document as a certification of hazard assessment.”

OSHA’s Bloodborne Pathogens standard requires that labs perform an exposure risk determination for each employee. Labs must assess exposure risk levels by job classification, and then assess exposure risk for tasks performed in the department. The Hazard Communication standard explains the need for chemical hazard determination. There are many types of risk assessments that must be performed, and regulations stipulate that they must be reviewed and updated (if necessary) every year. Things change in a dynamic department like the laboratory, and understanding the changing risks of harm can be key to keeping staff safe.

The four basic steps included in a risk assessment are hazard identification, identifying those at risk, choosing control measures, and reviewing the findings. It may sound easy, but lab hazards can come in many forms (physical, mental, chemical, biological, etc.), so walk around the department to look for those you may have missed. Review incident records as well to see what harm is occurring in the lab. Next, determine what employees may be harmed and how. Consider those who work in the lab each day and those that are just passing through the area. An evaluation of the risks follows. If the risk cannot be removed, decide what controls (engineering, administrative, PPE) need to be in place. Finally, review each risk assessment on a regular basis. Things change in the lab, and with change may come new hazard risks – or even the reduction of potential harm if risks have been reduced via elimination or substitution. Again, examine these assessments at least annually or whenever major changes occur in a particular lab safety arena.

For many laboratories, the advent of the COVID-19 pandemic brought new testing platforms and procedures to the department, and this testing had to be implemented quickly. Is there a way to use risk assessments to help introduce new processes safely? Absolutely! The use of a standard form to assess the potential hazards of new or updated processes and/or equipment is actually a high quality finishing touch on an overall assessment program, and unfortunately, it is something that is often missing in many labs.

Considering the tasks, sample handling, equipment, reagents, and overall biosafety of the new process, choose the likelihood of hazard incidents (rare, possible, likely, etc.), and classify the consequences of each occurrence (minor, moderate, major, etc.). Use a matrix to calculate the overall level of risk for the new procedure or equipment. For example, if a new COVID-19 test platform requires opening samples, the likelihood and risk of exposure may be designated as “high” or even “very high.” Next, determine the controls that should be put in place to decrease the exposure opportunities. For example, if centrifuge rotor covers, a biological safety cabinet, and a surgical mask is added to the normal lab PPE, the overall risk of that testing process is reduced. Document the decision and keep those records available for any future reviews that may be needed. Once the assessment is complete, complete the appraisal of the new process with a quick safety audit. Look for additional biohazards encountered, chemical safety, electrical safety, and even potential waste handling issues. If you couple that safety analysis with the risk assessment, you are doing an above-average job at circumventing hazards in the department. (For examples, go to https://www.aphl.org/programs/preparedness/Documents/APHL%20Risk%20Assessment%20Best%20Practices%20and%20Examples.pdf)

Keeping staff safe from exposures and injuries in the laboratory is a massive and time-consuming task, but it is required by our regulatory agencies and it needs to be a top priority. When used properly and completely, a risk assessments can be a powerful tools that begins your look into safety hazards and then closes the loop to avert them. Having an awareness and control of departmental hazards is one way to rock safety in the laboratory.

Dan Scungio, MT(ASCP), SLS, CQA (ASQ) has over 25 years experience as a certified medical technologist. Today he is the Laboratory Safety Officer for Sentara Healthcare, a system of seven hospitals and over 20 laboratories and draw sites in the Tidewater area of Virginia. He is also known as Dan the Lab Safety Man, a lab safety consultant, educator, and trainer.

Overview of Laboratory Tests for Cytomegalovirus


Cytomegalovirus (CMV) is considered the most important pathogen in transplant recipient patients as it can cause significant morbidity and mortality. Anti-CMV treatments have proven to be effective but are not without adverse side effects. Thus, there is a strong need for sensitive and reliable tests to diagnose and monitor active CMV infection. Several testing methodologies are available in today’s clinical laboratories to evaluate a patient’s CMV status: viral culture, serology, histopathology, pp65 antigenemia, and quantitative PCR. In this post, we will review the advantages and limitations of these tests.

Viral culture

Viral culture is performed most commonly by the shell vial assay (also known as rapid culture), in which a cell line (usually human fibroblast cells) is inoculated with patient sample by centrifugation. The virus is then detected by either direct or indirect fluorescent monoclonal antibody, providing results within 1-3 days. The centrifugation step greatly improves turnaround time when compared to traditional tube cell culture technique, which may take 2-3 weeks before a result can be reported as negative.

Culturing CMV has been largely replaced by newer methodologies like quantitative PCR and CMV antigenemia. This is due to relatively weaker sensitivity for diagnosing CMV infection compared to newer tests, as well as slower turnaround time. Viral cultures of urine, oral secretions, and stool are not recommended due to poor specificity; however, for diagnosis of congenital CMV, viral culture of urine or saliva samples is an acceptable alternative if PCR is not available.


CMV serostatus is an important metric to evaluate prior to patients receiving a hematopoietic or solid organ transplant. Serologic testing is done primarily via enzyme immunoassays and indirect immunofluorescence assays. These tests check for presence of anti-CMV immunoglobulin (Ig)M and IgG to provide evidence of recent or past infection. Outside of establishing serostatus (primarily in organ donors and recipients), serologic testing for CMV is not recommended in diagnosing or monitoring active CMV infection.

CMV IgM antibodies can be detected within the first two weeks of symptom development and can be present for another 4-6 months. IgG antibodies can be detected 2-3 weeks after symptoms develop, and remain present lifelong. These antibody measurements are particularly useful in determining risk of CMV acquisition in seronegative patients (negative for IgM and IgG) at time of transplantation. IgG titers can also be measured every 2-4 weeks to assess for CMV reactivation in seropositive patients. Since CMV IgG persistently remains in circulation, testing for it has a higher specificity compared to IgM, and thus is the preferred immunoglobulin to test for in establishing serostatus. Serologic tests can be falsely positive if patients have recently received IVIG or blood products, so testing on pretransfusion samples are preferred if possible.


Under the microscope, cells infected with CMV can express certain viral cytopathic effects. These infected cells classically show cytoplasmic and nuclear inclusions (owl eye nuclei) with cytoplasmic and nuclear enlargement. Additionally, immunohistochemistry (IHC) can stain antibodies specifically for CMV proteins to highlight infected cells, making histologic examination quicker and improving diagnostic sensitivity.

Histopathology can be useful in identifying tissue-invasive disease, such as CMV colitis or pneumonitis. Cases in which PCR testing is negative does not necessarily exclude tissue-invasive disease; thus, the diagnosis of tissue-invasive disease relies on histologic examination (with or without IHC) or possibly viral culture. On the other hand, a negative histologic result does not exclude tissue-invasive disease, possibly due to inadequate sampling, and shows the potential for weak diagnostic sensitivity.

pp65 antigenemia

CMV antigenemia testing uses indirect immunofluorescence to identify pp65 antigen, a CMV-specific matrix protein, in peripheral blood polymorphonuclear leukocytes. Whole blood specimens are lysed and then the leukocytes are cytocentrifuged onto a glass slide. Monoclonal antibodies to pp65 are applied, followed by a secondary FITC-labeled antibody. The slide is then read using a fluorescence microscope for homogenous yellow-green polylobate nuclear staining, indicating presence of CMV antigen-positive leukocytes. Studies have suggested that a higher number of positive cells correlates with an increased risk of developing active disease. The sensitivity of antigenemia testing is higher than that of viral culture and offers a turnaround time within several hours.

This test has been utilized since the 1980s, but has seen less use recently due to the increasing popularity of quantitative PCR. Antigenemia testing is labor intensive, and requires experienced and trained personnel to interpret the results (which can be somewhat subjective). This test also must be performed on whole blood specimens within 6-8 hours of collection due to decreasing sensitivity over time, which limits transportability of specimens. Additionally, It is not recommended to be run on patients with absolute neutrophil counts below 1000/mm3, due to decreased sensitivity. Despite these limitations, CMV antigenemia testing is still considered a viable choice for diagnosing and monitoring CMV infection, especially when viral load testing is not available.

Quantitative PCR

Quantitative real-team polymerase chain reaction (PCR) is the most commonly used method to monitor patients at risk for CMV disease and response to therapy, as well as for diagnosing active CMV infection. The advantages of using a quantitative PCR assay include increased sensitivity over antigenemia testing, quick turnaround time, flexibility of using whole blood or plasma specimens for up to 3-4 days at room temperature, and the availability of an international reference standard published by the World Health Organization (WHO).

Several assays from Roche, Abbott, and Qiagen are available and FDA-approved. The targets of these assays vary, with some targeting polymerase and other targeting CMV major immediate early gene. These assays are all calibrated with the WHO international standard, which was developed in 2010 to help standardize viral load values among different labs when results are reported in international units/mL. The goal of this international standard is to decrease the interlaboratory variability of viral load, and determine the appropriate cut-offs for determining clinical CMV disease. There is still improvement to be made in this area, as variability still exists between labs.


There are several tests to determine the CMV status of patients. Some of these tests are suited for particular goals, such as serology for determining serostatus prior to organ transplantation, or histology and IHC to diagnose tissue-specific CMV disease. For diagnosis and monitoring of general CMV disease, the test of choice in most laboratories is quantitative PCR, which offers automated, quick and sensitive results. Antigenemia, while dated and labor intensive, is still an acceptable alternative when PCR is neither available nor cost-effective for smaller labs. Both of these testing methods are preferred over viral culture, which has poorer diagnostic sensitivity and relatively longer turnaround time.

Despite the numerous advantages quantitative PCR has, there is still variability in viral load counts among laboratories. This is due to varying DNA extraction techniques, gene targets used by PCR, and specimen types used. There is still a lot of work to be done in standardizing testing in regards to not just CMV, but also other viral pathogens like Epstein-Barr virus, BK virus, adenovirus and HHV6. Updated standards and increased use of standardized assays will hopefully decrease this variability between labs to improve testing results and in turn, improve patient care.


  1. https://www.uptodate.com/contents/overview-of-diagnostic-tests-for-cytomegalovirus-infection#H104411749
  2. https://www.uptodate.com/contents/congenital-cytomegalovirus-infection-clinical-features-and-diagnosis?topicRef=8305&source=related_link#H9542666
  3. Kotton CN, Kumar D, Caliendo AM, et al. Updated international consensus guidelines on the management of cytomegalovirus in solid-organ transplantation. Transplantation. 2013;96(4):333-60.
  4. Hayden RT, Sun Y, Tang L, et al. Progress in Quantitative Viral Load Testing: Variability and Impact of the WHO Quantitative International Standards. J Clin Microbiol. 2017;55(2):423-430.
  5. Kotton CN, Kumar D, Caliendo AM, et al. The Third International Consensus Guidelines on the Management of Cytomegalovirus in Solid-organ Transplantation. Transplantation. 2018;102(6):900-931.

-David Joseph, MD is a 2nd year anatomic and clinical pathology resident at Houston Methodist Hospital in Houston, TX. He is planning on pursuing a fellowship in forensic pathology after completing residency. His interests outside of work include photography, playing bass guitar and video games, making (and eating) homemade ice cream, and biking.