Microbiology Case Study: Blood stream infection in a 77 year old patient – is it Really from Mosquitoes?

A 77 year old male with a past medical history of end stage renal disease (ESRD) on hemodialysis, type 2 diabetes, coronary and peripheral artery disease, and squamous cell carcinoma of the lung on current chemotherapy/radiation was admitted to our hospital from his outpatient hematology oncology clinic for acute hypoxia. Due to an episode of decreased responsiveness and a potential stroke, Head computed tomography (CT) and computed tomography angiography (CTA) were performed. Electroencephalography showed diffuse slowing, suggestive of encephalopathy. Three days after admission, he became hypotensive and febrile. Pulmonology/critical care was consulted; blood and respiratory samples collected for cultures. The blood culture grew gram negative rods in the aerobic bottles (Images 1-3) after overnight incubation. The patient was initially on cefepime and switched to meropenem 500 mg IV daily.

The following day, the blood culture isolate was identified as Elizabethkingia anophelis. The isolate was resistant to both of the patient’s prior inpatient antibiotics, cefepime and meropenem. Additionally, the isolate was resistant to first, second, and third generations of cephalosporins, aztreonam, tetracyclines and tobramycin. However, it was susceptible to amikacin, ciprofloxacin, gentamicin, and trimethoprim/sulfamethoxazole. Meropenem was discontinued and replaced with ciprofloxacin 400 mg IV daily. Infectious disease was consulted; at this time the patient was displaying nuchal rigidity and extreme encephalopathy. Increased dosing of Ciprofloxacin for better central nervous system penetration, in combination with trimethoprim/sulfamethoxazole 2.5mg/kg IV q8h, rifampin 600 mg IV daily was recommended and a lumbar puncture to be performed once the patient was stable. Sadly, due to underlying severe comorbidities, along with worsening CNS responses, the patient expired on day 9.

Image 1. Small gram negative rods of E. anophelis from positive aerobic blood cultures.
Image 2. Blood Agar Plate growing E. anophelis after overnight incubation at 35 degrees C
Image 3: Chocolate plate growing smooth creamy gram negative E. anophelis after overnight incubation at 35 degrees C


The genus Elizabethkingia was named after Elizabeth O. King, a microbiologist at the Center for Disease Control (CDC) and Prevention, who discovered many medically important bacteria in the late 1940s to early 1960s. This included describing Elizabethkingia meningoseptica (formerly Chryseobacterium meningosepticum) in 1959. Elizabethkingia and Kingella genera, and the species Kingella kingae are also named in her honor.1 Elizabethkingia is a gram negative, obligate aerobic bacillus. It was classified under the families Flavobacteriaceae and Chryseobacterium, but was reclassified as Elizabethkingia in 2005.2 E. meningoseptica, the most frequently isolated Elizabethkingia species, has been implicated in cases of neonatal sepsis, meningitis, and nosocomial pneumonia.3

On the other hand, E. anophelis was recently characterized in 2011 and was initially thought to be underrepresented likely due to the genotypic and phenotypic similarity to E. meningoseptica.4,5,6 The species name Elizabethkingia anophelis originated from the anopheles mosquito as it has been isolated from the midgut of Anopheles gambiae mosquitoes.4 The role of mosquitos in maintenance and transmission of Elizabethkingia anophelis is unclear.4,6 Oxidase and catalase positive E. anophelis, (Fig 1) grows well on blood and chocolate agar plates (Fig 2 and 3) as smooth and slight-yellowish colonies although it does not grow on MacConkey agar.7

Beginning late 2015, an increased number of Elizabethkingia infections were identified in Southeastern Wisconsin. Between November 2015 to May 2016, 63 cases of E. anophelis were reported to the Wisconsin Division of Public Health. Cases spread across Illinois and Michigan as well, making it the largest E. anophelis outbreak described to date. A case series published from Froedtert Health System hospitals described their experience with E. anopheles.8 This was a retrospective case series of all consecutive patients admitted to Froedtert Health System hospitals with positive cultures of any site for Elizabethkingia, Flavobacterium, and Chryseobacterium from November 2015 to June 2016. In this time period, 11 patients were identified with cultures positive for E. anophelis. All patients had positive blood cultures at the time of hospital admission. E. anophelis was identified in both sterile and nonsterile body fluids. All 11 patients had at least one major comorbidity, including cancer, COPD, diabetes, ESRD requiring hemodialysis, and alcohol abuse. Two patients died within 30 days of a positive E. anopheles culture (attributable mortality rate, 2/11 = 18.2%).5, 8, 9

Interestingly, vertical transmission of E. anopheles causing neonatal meningitis has been reported.6 Molecular evidence suggested vertical transmission from a mother with chorioamnionitis, but a mechanism of colonization for the mother could not be found and environmental contamination was not also found.6,7 On the other hand, taps and aerators contaminated with E. anophelis in an intensive care unit has been reported.10 E. anophelis should be treated as a true pathogen, particularly in patients with multiple comorbidities.8 Isolation in sterile fluid should never be considered a contaminant.

Since Elizabethkingia is a non-glucose, non-lactose-fermenter, the MIC breakpoint of E. anophelis is reported based on those of non-Enterobacterales Table 2B-5 of CLSI (Clinical Laboratory Standard Institute) M100 guidelines. Elizabethkingia species, including E. anophelis, are intrinsically resistant to several antibiotics and produce elevated MIC on in vitro susceptibility tests. A number of species also harbor beta-lactamase/metallo beta-lactamase (MBL) genes. Empirical treatment should include piperacillin/tazobactam plus quinolone, rifampin, or minocycline. Vancomycin has been used in severe infections, especially meningitis. The best duration of therapy has not been evaluated by clinical trials8.

In summary, our patient acquired this infection in the setting of multiple chronic comorbidities and was immunocompromised due to active malignancy and recent chemotherapy. He has a similar clinical profile to the other patients in the above-mentioned study. One notable difference is that our patient’s isolate was resistant to cefepime, where the isolates from this outbreak were susceptible. After discussion with our infectious disease colleagues regarding this case, we agreed his cause of death was likely multifactorial, though this infection may have been a significant contributing factor.


  1. KING EO. Studies on a group of previously unclassified bacteria associated with meningitis in infants. Am J Clin Pathol. 1959 Mar;31(3):241-7. doi: 10.1093/ajcp/31.3.241. PMID: 13637033.
  2. Kim KK, Kim MK, Lim JH, Park HY, Lee ST. Transfer of Chryseobacterium meningosepticum and Chryseobacterium miricola to Elizabethkingia gen. nov. as Elizabethkingia meningoseptica comb. nov. and Elizabethkingia miricola comb. nov. Int J Syst Evol Microbiol. 2005 May;55(Pt 3):1287-1293. doi: 10.1099/ijs.0.63541-0. PMID: 15879269.
  3. Jean SS, Lee WS, Chen FL, et al. Elizabethkingia meningoseptica: an important emerging pathogen causing healthcare-associated infections. J Hosp Infect 2014; 86:244–9.
  4. Kämpfer P, Matthews H, Glaeser SP et al. . Elizabethkingia anophelis sp. nov., isolated from the midgut of the mosquito Anopheles gambiae. Int J Syst Evol Microbiol 2011; 61(Pt 11):2670–5. [PubMed] [Google Scholar]
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  6. Lau, Susanna K.P.; Wu, Alan K.L.; Teng, Jade L.L.; Tse, Herman; Curreem, Shirly O.T.; Tsui, Stephen K.W.; et al. (February 2015). “Evidence for Elizabethkingia anophelis Transmission from Mother to Infant, Hong Kong”. Emerging Infectious Diseases. 21 (2): 232–241. doi:10.3201/eid2102.140623. PMC4313635. PMID25625669
  7. Koneman’s Color Atlas and Textbook of Diagnostic Microbiology. 7th Edition. 2016.
  8. Castro, C. E., Johnson, C., Williams, M., Vanderslik, A., Graham, M. B., Letzer, D., . . . Munoz-Price, L. S. (2017). Elizabethkingia anophelis: Clinical Experience of an Academic Health System in Southeastern Wisconsin. Open Forum Infectious Diseases, 4(4). doi:10.1093/ofid/ofx251
  9. Wisconsin Department of Health Services; Elizabethkingia 2017. Available at: https://www.dhs.wisconsin.gov/disease/elizabethkingia.htm. Accessed 9 January 2017. [Google Scholar]
  10. Balm MN, Salmon S, Jureen R, Teo C, Mahdi R, Seetoh T, Teo JT, Lin RT, Fisher DA. Bad design, bad practices, bad bugs: frustrations in controlling an outbreak of Elizabethkingia meningoseptica in intensive care units. J Hosp Infect. 2013 Oct;85(2):134-40. doi: 10.1016/j.jhin.2013.05.012. Epub 2013 Aug 17. PMID: 23958153.

-J. Stephen Stalls, MD is a PGY-II pathology resident at the East Carolina University Department of Pathology and Laboratory Medicine. He plans to pursue hematopathology and molecular pathology fellowships, but also greatly enjoys his time in the microbiology lab. Outside of work, he enjoys playing the drums and going to concerts.

-Phyu Thwe, Ph.D., D(ABMM), MLS (ASCP)CM is a Technical Director at Vidant Medical Center Clinical Microbiology Laboratory. She completed a Clinical and Public Health Microbiology Fellowship through a CPEP-accredited program at the University of Texas Medical Branch (UTMB) in Galveston, Texas. She is interested in extrapulmonary tuberculosis and developing diagnostic algorithms.

Breakpoint Breakdown II: Breakpoints at the Bedside

A critical task for clinical microbiologists is interpreting antimicrobial susceptibility testing (AST) results for microorganisms recovered from patient specimens. But what happens to that information after it is passed along to the clinical team, and how does it influence patient care at the bedside? In our previous blog, we discussed details concerning how breakpoints are established and identified them as an indispensable component of appropriate and effective antimicrobial prescribing. Here we will discuss the more practical application of breakpoints to guide patient care. To aid in the discussion, I’ve once again recruited two infectious diseases (ID) pharmacists from the UT Southwestern Medical Center to provide their valuable prospective on the use of AST results both at the bedside and to optimize antimicrobial stewardship initiatives.

Applying Breakpoints in Clinical Care

In the most simplistic terms, the reporting of isolate’s categorical susceptibility (susceptible, intermediate, and resistant) provides guidance concerning which antimicrobials will likely be effective to treat an infection.1 Many microbiology laboratories, however, report additional AST information including minimal inhibitory concentrations (MICs) and phenotypic information concerning resistance mechanisms (e.g. production of Extended Spectrum Beta-Lactamases (ESBLs) or inducible-clindamycin resistance)2. This more detailed information is often utilized by ID specialists to help guide and optimize beside management.

Unfortunately, several misapplications of AST data can be encountered in clinical care (Image 1). One common error is simply selecting the antimicrobial with the lowest MIC for treatment. This runs contrary to the idea that each antimicrobial has an individualized breakpoint for a given pathogen or pathogen group. Therefore, a low MIC for antimicrobial “A” may be at or near the breakpoint, but higher MIC for antimicrobial “B” may be a dilution or more lower than the breakpoint. Another common misconception is that if the isolate is reported susceptible to a given antimicrobial, that antimicrobial is the optimal choice for the patient, and will work 100% of the time! Very little in medicine is 100%, and this holds true for antimicrobials. Even if the optimal agent is selected based on AST data, there is no guarantee it will be effective in the patient. Many factors influence the effectiveness of antimicrobials including patient characteristics (e.g., immune system, kidney function), drug characteristics (i.e., pharmacokinetics [PK]/pharmacodynamics[PD]), and bug characteristics (e.g., underlying resistance present despite reporting as susceptible or potential for inducible resistance).2 This illustrates why clinicians may prefer to have all available information to further aid in optimization of antimicrobial therapy beyond the information provided by categorical reporting.

But why is an MIC needed – isn’t just knowing something is susceptible or resistant enough? In particular, more granular information such as the actual MIC for a given bug-drug combination may aid in more intricate decisions surrounding the choice of agent and dose. One argument for routine reporting of MIC values is the ability to better discern how near the breakpoint the particular MIC is for a given drug. The standard MIC reporting error is plus or minus one doubling dilution. When this is factored in, it may provide additional context as to how near or over the isolates phenotype is to assigned breakpoint.2 Clinicians may further couple this with their understanding of PK/PD to determine the optimal agent based on antimicrobial properties and microbial characteristics.

A counter argument is that the MIC values are prone to variation depending on testing modality and may not represent truly what occurs in actual humans (see Mouton JW et al. for more detailed discussion).3 As mentioned above, some argue that MIC reporting leads to misinterpretations of AST by a provider’s tendency to pick the lowest number, and thus prefer categorical reporting for simplification. Provider education and selective or cascade reporting are two strategies that may help clinicians select the optimal agent, but also have inherent limitations. All this taken into consideration, we feel the benefit of antimicrobial optimization supports reporting the MIC.

Applying Breakpoints at the Bedside

Breakpoints and MIC data are utilized at the bedside to avoid utilization of suboptimal agents due to issues with the drug and bug. For example, underlying resistance patterns may not be overtly apparent to clinicians. Depending on an institution’s reporting criteria, an E. coli isolate may be reported as susceptible to piperacillin/tazobactam despite being identified as an ESBL producer. In this instance, piperacillin/tazobactam may be suboptimal for certain infections based on available outcomes data, with some supposition that reported MIC values may be responsible for improper drug selection and treatment failures.4 Therefore, clinicians may favor alternative agents for infections despite documented susceptibility. This example also illustrates that certain resistance patterns (e.g., ceftriaxone resistance as surrogate for possible ESBL production) may help alert clinicians to underlying resistance, steering them clear of potential suboptimal agents reported as susceptible.

ID pharmacists often utilize MIC values to determine the optimal agent and dose for a given patient based on a working knowledge of breakpoint relationships and PK/PD. This may be taken a step further in the setting of an infection with a multidrug resistant organism which exhibit MICs in the intermediate or susceptible dose-dependent range. For certain drug-bug combinations with elevated MICs, it may be possible for the ID pharmacist to devise an optimized and personalized therapeutic regimen that will likely overcome the elevated MIC, either as monotherapy or utilizing potential synergistic combinations of different antibiotics.

The application of dose-dependent breakpoints, as seen with daptomycin for Enterococcus faecium, help illustrate the importance of understanding and applying dosing concepts in the context of established clinical breakpoints.1 This allows the clinician to not only optimize dosing, but also understand that an MIC at or near the breakpoint may warrant combination therapy in the setting of a serious infection. A lack of understanding of the relationship between breakpoints and dosing could potentially lead to an inadvertent assumption that lower FDA-approved doses would be reasonable despite the breakpoint being based on higher dosing strategies. These are just a small sampling of the clinical applications utilized on a daily basis to optimize a particular patient’s antimicrobial therapy.

A more global application of breakpoints occurs within antimicrobial stewardship programs. Antimicrobial stewardship focuses on the appropriate and optimal use of antimicrobials. To aid in this goal, AST reporting is key to developing clinical guidance on treating a variety of infections. Most notably, the optimization of empiric antimicrobial choices is aided by local or institutional susceptibility patterns. The cumulative antibiogram is pivotal to establishing optimal empiric antimicrobial choices across the spectrum of infectious diseases.5 Importantly, breakpoints are revised when new or additional clinical information becomes available, further illustrating the need to understand how a given institution implements and applies them. There is always the potential that from one year to the next susceptibilities may drop substantially due to implementation of new breakpoints, not necessarily an increased incidence of pathogens with a certain resistance pattern.6 In addition, the ability to adopt CLSI breakpoints often lags well beyond the release of the updated breakpoints, necessitating clinicians to be cognizant of changes and carefully assess categorical reporting as it may be discordant with the most up-to-date breakpoints.1,6 This topic will be the focus of our next blog.

More specific application of AST data can be made for either site (e.g., blood, urine, etc.), infection type (e.g., community-acquired pneumonia), or organism-specific MIC distributions. A recent trend in national guideline recommendations centers on ascertaining local susceptibility and pathogen distribution to provide optimal antimicrobial choices for certain infections (e.g., pneumonia).7 In addition, focused review of individual pathogen MIC distributions provide another level of detail that helps better define upfront institutional antimicrobial dosing strategies to optimize PK/PD parameters. In conclusion, the reporting of AST data remains a cornerstone for the effective management of infections. Information collected by laboratory technicians provides invaluable information to clinicians. This information helps optimize the selection and dosing of antimicrobial agents at the bedside, which in turn improves the clinical management of patients and infection-related outcomes.

Image 1: Clinical scenario outlining the application of MIC and breakpoint data at the bedside.


  1. Clinical and Laboratory Standards Institute. Development of In Vitro Susceptibility Testing Criteria and Quality Control Parameters, 5th Edition (M23).
  2. Giuliano C, Patel CR, and Kale-Pradhan PB. A Guide to Bacterial Culture Identification and Results Interpretation. P&T. 2019 April 44(4):192-200.
  3. Mouton JW, Muller AE, Canton R, et al. MIC-based dose adjustment: facts and fables. J Antimicrob Chemother 2018; 73:567-68.
  4. Pogue JM and Heil EL. Laces out Dan! The role of tazobactam based combinations for invasive ESBL infections in a post-MERINO world. E Expert Opin Pharmacother.2019 Dec;20(17):2053-57.
  5. Clinical and Laboratory Standards Institute. Analysis and Presentation of Cumulative Antimicrobial Susceptibility Test Data; Approved Guideline, 4th Edition (M39-A4).
  6. Humphries RM, Abbott A, Hindler JA. Understanding and Addressing CLSI Breakpoint Revisions: a Primer for Clinical Laboratories. J Clin Microbiol. 2019 May 24;57(6):e00203-19.
  7. Khalil AC, Metersky ML, Klompas M, et al. Management of Adults with Hospital-acquired and Ventilator-associated Pneumonia: 2016 Clinical Practice Guidelines by the Infectious Diseases Society of America and the American Thoracic Society. Clin Infect Dis. 2016 Sep 1;63(5):e61-e111.

-James Sanders, PharmD, PhD, is an infectious diseases pharmacy specialist and assistant professor at the University of Texas Southwestern Medical Center in Dallas, Texas. His academic and research interests are focused on multi-drug resistant Gram-negative bacteria, surgical site infections and HIV pharmacotherapy.

-Marguerite Monogue, PharmD is an infectious diseases pharmacy specialist and assistant professor at the University of Texas Southwestern Medical Center in Dallas, Texas. She is interested in antimicrobial pharmacokinetics/pharmacodynamics and multi-drug resistant Gram-negative bacteria.

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

Microbiology Case Study: A 40 Year Old Man with LVAD Exit Site Pain

Case history

A 40 year old male with a history of cardiomyopathy requiring a left ventricular assist device (LVAD) was seen in clinic with a complaint of pain at the exit site of the LVAD driveline. History is notable for multiple admissions for driveline-associated complications. Despite extensive prior evaluation, cultures and imaging of the driveline exit site had been repeatedly negative with the exception of a methicillin-susceptible Staphylococcus aureus.This was treated with nafcillin, followed by doxycycline for oral suppression. The patient had stopped taking oral antibiotics two months prior to presentation. Imaging revealed a 1.4 cm region around the driveline exit site suggestive of either phlegmon, hematoma, or a developing abscess. Blood cultures and cultures of the driveline exit site were collected and sent to the clinical microbiology laboratory. Upon physical examination, the driveline exit site was tender, but no erythema was noted. The patient endorsed intermittent rust-colored drainage from the site. Blood cultures remained negative for the duration of the patient’s hospital course, and the patient was discharged on nafcillin with plans to transition to doxycycline.

Laboratory identification

The Gram stain of the driveline exit site was unremarkable, with no organisms and few neutrophils seen. Aerobic cultures yielded a light amount of gram positive cocci in addition to mixed skin flora. Colonies were small, and weakly beta hemolytic on blood agar (Image 1A). This organism was catalase- and coagulase-positive, and definitively identified as Staphylococcus aureus by MALDI-TOF MS. Susceptibility testing was performed by broth microdilution, where the organism was determined to be a vancomycin-intermediate Staphylococcus aureus (VISA, MIC=4, Image 1C). Due to the unusual nature of the result, it repeated and confirmed by E-test (Image IB) in our laboratory, and independently verified at our contract reference laboratory. The isolate was also referred to the Texas State Public Health Laboratory where the vancomycin-intermediate phenotype was again confirmed. This isolate was also daptomycin non-susceptible, but remained susceptible to oxacillin, trimethoprim/sulfamethoxazole, linezolid, rifampin, and clindamycin.

Image 1. A. Growth of the weakly beta-hemolytic vancomycin-intermediate S. aureus strain on blood agar. B. Measurement of vancomycin susceptibility by E-test (MIC=4). C. Confirmation of the VISA phenotype by broth microdilution (MIC=4).


All models of LVADs require a percutaneous driveline which is a link between the implanted device and the external power source.1In addition to providing power, the driveline also provides controlling and sensing functions for the LVAD.2 The driveline exit site is one of the most common sites of LVAD infection as the driveline creates a conduit for entry of bacteria from the external environment. Additionally, the prosthetic material of the driveline can serve as an ideal substrate for biofilms formation.1 The most common microorganisms associated with LVAD-related infections members of the skin microbiota (i.e. staphylococci), Pseudomonas sp., and enteric bacteria.3

Staphylococcus aureus remains an important human pathogen globally. While antibiotic intervention remains a mainstay of treatment, the emergence of resistance has historically changed the way patients are managed. Mobile genetic elements (including plasmids and transposons) are important mediators of antibiotic resistance in S. aureus, particularly with respect to beta-lactams and glycopeptide antibiotics. Due to the widespread emergence of beta-lactamase conferred penicillin-resistance, semisynthetic penicillinase-resistant penicillins (including methicillin, oxacillin, and nafcillin) were developed for clinical use in the late 1950s. However, resistance to these compounds in S. aureus was reported only a few years following their introduction. Vancomycin became the antibiotic of choice for methicillin-resistant S. aureus (MRSA) therapy in the 1980s, and contemporary management remains largely reliant on this antibiotic despite the recent availability of newer agents from different antibiotic classes.4Thus, vancomycin non-susceptibility among S. aureus isolates is a rare phenomenon with serious clinical implications, with only modest increases in vancomycin MICs resulting in treatment failures.5

The first vancomycin-intermediate S. aureus (VISA) isolate was reported in 1997 in Japan, followed by the first vancomycin-resistant isolate in 2002 in the US.4 It is important to note that the mechanisms driving these two phenotypes are entirely different. The fully vancomycin-resistant phenotype is due to the acquisition of the vanA gene which confers cell wall alterations that prohibit vancomycin from efficiently binding its target. By contrast, the vancomycin-intermediate phenotype remains less well described mechanistically, but VISA strains share similar phenotypic traits. These include: alterations in growth kinetics, increased cell wall thickness, a reduction in peptidoglycan crosslinking, decreased autolysis, altered surface protein profile, and variation of expression levels of global genetic regulators.4,5 These phenotypes are due to mutations and alterations in expression of a number of candidate genes involved in cell wall synthesis, capsule production, and global regulators of virulence.

The emergence of a VISA phenotype is usually found in the setting of MRSA strains that have been treated with prolonged vancomycin therapy.5 However, in this patient’s case, vancomycin had only been utilized infrequently for unrelated infections several years prior. Daptomycin had not previously been used in this patient’s clinical care. This VISA isolate was also oxacillin-susceptible which is a less common finding among reported VISA strains. While exposure of S. aureus to non-glycopeptide antibiotics including beta-lactams can trigger VISA phenotypes in vitro,6 it is currently not possible to elucidate the mechanism underpinning vancomycin non-susceptibility, nor what has driven this resistant phenotype, in this patient’s isolate. The patient currently is doing well on doxycycline suppressive therapy after completing his course of nafcillin, and continues to be monitored through follow-up appointments.


  1. Leuck A-M. 2015. Left ventricular assist device driveline infections: recent advances and future goals. Journal of Thoracic Disease 7:2151-2157.
  2. Long B, Robertson J, Koyfman A, Brady W. 2019. Left ventricular assist devices and their complications: A review for emergency clinicians. The American Journal of Emergency Medicine 37:1562-1570.
  3. Zinoviev R, Lippincott CK, Keller SC, Gilotra NA. 2020. In Full Flow: Left Ventricular Assist Device Infections in the Modern Era. Open Forum Infectious Diseases 7.
  4. McGuinness WA, Malachowa N, DeLeo FR. 2017. Vancomycin Resistance in Staphylococcus aureus
The Yale Journal of Biology and Medicine 90:269-281.
  5. Gardete S, Tomasz A. 2014. Mechanisms of vancomycin resistance in Staphylococcus aureus. The Journal of Clinical Investigation 124:2836-2840.
  6. Roch M, Clair P, Renzoni A, Reverdy M-E, Dauwalder O, Bes M, Martra A, Freydière A-M, Laurent F, Reix P, Dumitrescu O, Vandenesch F. 2014. Exposure of Staphylococcus aureus to subinhibitory concentrations of β-lactam antibiotics induces heterogeneous vancomycin-intermediate Staphylococcus aureus. Antimicrobial agents and chemotherapy 58:5306-5314.

-Zoya Khan MS, MLS(ASCP)CM is a medical technologist in the clinical microbiology laboratory at UT Southwestern with almost 10 years’ experience. She received a BS in Medical Technology from Texas Women’s University, and an MS in Clinical Practice Management from Texas Tech Health Science Center. She has an active interest in mycology and laboratory assay verification.

Francesca Lee, MD, is an associate professor in the Departments of Pathology and Internal Medicine (Infectious Diseases) at UT Southwestern Medical Center.

-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.

A Cording Too: “Cording” in Clinical Isolates of Mycobacterium

A cornerstone of good clinical microbiology laboratory practice is to look for visual clues in how organisms grow in culture. This can help quickly signal to the laboratorian that a particularly meaningful pathogen is present.

For example, a wound culture where an anaerobic blood agar plate is showing a double zone of hemolysis? The tech should immediately think that Clostridium perfrigens may be present. An abscess culture growing a ridiculously mucoid colony of a lactose-fermenting Gram negative rod? Hyper-mucoid Klebsiella pneumoniae is a hypervirulent strain associated with abscess formation. Training and experience in what to look for in cultures is one of the fascinating and exciting (and potentially daunting) aspects of clinical microbiology!

But crucially, sometimes the textbook visual association is NOT as specific a finding as we may believe! One such example: Mycobacterium species and cording.

Cording and M. tuberculosis

“Cording” is term used to describe twisting, serpentine appearance of Mycobacterium in liquid broth culture. And, at least in many places, it is taught to budding microbiologists and infectious disease clinicians as a hallmark characteristic of Mycobacterium tuberculosis (Mtb).

How long have people been observing Mtb cording in liquid culture? From very beginning!

Robert Koch, originator of the famous Koch’s Postulates, used Mtb (now recognized to be just one member of the M. tuberculosis complex) to demonstrate that disease was caused by discrete organisms originating from one host and infecting another, what we accept as germ theory. In his original description of Mtb, Koch wrote that the bacilli would “ordinarily form small groups of cells which are pressed together and arranged in bundles.”

Cording phenotype is distinct from “clumping.” With cording, bacilli lay in tightly packed parallel strands, and not clumped together showing random bacilli orientation.

The phenomenon of cording is distinct to organisms cultured in liquid media like Middlebrook 7H9 broth. But in 1947 Middlebrook (where Middlebrook 7H9 broth gets its name), Dubos and Pierce published a key paper showing that cording Mtb strains also grew as rough colonies on solid agar and that this phenotype was associated with virulent and not avirulent Mtb strains.

Figures from 1947 Middlebrook et al., showing avirulent Mtb clumping (1a) and smooth colonies (2a) and virulent Mtb with cording (1b) and rough colonies (2b).

Only a few years later in 1950, the actual cell wall component, the so-called “cording factor,” that causes these growth characteristics was extracted and identified. Trehalose-6,6-dimycolate (TDM), a major glycolipid from the cell walls of virulent strains of Mtb.

Further work suggested that, not only was this TDM glycolipid the cause of the unique cording appearance and rough colonies in in vitro culture conditions, but that directly was a virulence factor: studies in animal models showed that TDM not only directly allowed Mtb to avoid phagocytosis from macrophages by virtue of forming large clusters of bacilli, but it also directly prevented macrophage intracellular killing mechanisms (see Hunter, 2006).

The association between cording and clinical isolates of Mtb was seen as a very sensitive and specific finding. Such a robust feature, in fact, that a number of publications from clinical microbiology labs specifically to cording as a reliable method to quickly identify Mtb (see McCarter 1998) or at least to select which isolates should be further identified by more specific methods(see Nelson, 1998).

Cording in non-tuberculosis Mycobacteria

Though there was a strong the association between cording and Mtb, scientists had known for years that other Mycobacterium species, outside the M. tuberculosis complex, also expressed TDM. But beginning with a case report in 2008, an isolate of Mycobacterium marinum was seen to have a biofilm with “cording morphology.” Other reports of cording in M. marinum followed (see Staropoli 2008).

Importantly, cording in non-tuberculosis Mycobacterium species is 1) actual cording and 2) visually indistinguishable from cording seen in Mtb.

In my laboratory, (Spokane, Washington, USA) we regularly see isolates of Mycobacterium abscessus, a rapid growing mycobacterium species, with clear cording morphology. These twisting structure are clearly cords, not just clumps.

Researchers have since identified not just different levels of TDM in isolates of M. abscessus but also that those levels correlate to the known rough or smooth appearance of these isolates (see Llorens-Fons, 2017). It is interesting to consider that more virulent isolates of M. abscessus are likely to be rough, cording-type growers, similar to what is seen with Mtb.

Beyond what I hope was an interesting look into the history of Mycobacterium identification, and a great chance to show some beautiful AFB cording, it’s a good reminder that in our lab culture (pun definitely intended!) visual clues to organism identification may not fit what you were first taught.


  1. Middlebrook G, Dubos RJ, Pierce C. VIRULENCE AND MORPHOLOGICAL CHARACTERISTICS OF MAMMALIAN TUBERCLE BACILLI. J Exp Med. 1947 Jul 31;86(2):175-84. doi: 10.1084/jem.86.2.175. PMID: 19871665; PMCID: PMC2135722.
  2. Hunter RL, Olsen MR, Jagannath C, Actor JK. Multiple roles of cord factor in the pathogenesis of primary, secondary, and cavitary tuberculosis, including a revised description of the pathology of secondary disease. Ann Clin Lab Sci. 2006 Autumn;36(4):371-86. PMID: 17127724.
  3. McCarter YS, Ratkiewicz IN, Robinson A. Cord formation in BACTEC medium is a reliable, rapid method for presumptive identification of Mycobacterium tuberculosis complex. J Clin Microbiol. 1998 Sep;36(9):2769-71. doi: 10.1128/JCM.36.9.2769-2771.1998. PMID: 9705435; PMCID: PMC105205.
  4. Nelson SM, Cartwright CP. Comparison of algorithms for selective use of nucleic-acid probes for identification of Mycobacterium tuberculosis from BACTEC 12B bottles. Diagn Microbiol Infect Dis. 1998 Aug;31(4):537-41. doi: 10.1016/s0732-8893(98)00049-2. PMID: 9764392.
  5. Hall-Stoodley L, Brun OS, Polshyna G, Barker LP. Mycobacterium marinum biofilm formation reveals cording morphology. FEMS Microbiol Lett. 2006 Apr;257(1):43-9. doi: 10.1111/j.1574-6968.2006.00143.x. PMID: 16553830.
  6. Staropoli JF, Branda JA. Cord formation in a clinical isolate of Mycobacterium marinum. J Clin Microbiol. 2008 Aug;46(8):2814-6. doi: 10.1128/JCM.00197-08. Epub 2008 Jun 25. PMID: 18579723; PMCID: PMC2519507.
  7. Llorens-Fons M, Pérez-Trujillo M, Julián E, Brambilla C, Alcaide F, Byrd TF, Luquin M. Trehalose Polyphleates, External Cell Wall Lipids in Mycobacterium abscessus, Are Associated with the Formation of Clumps with Cording Morphology, Which Have Been Associated with Virulence. Front Microbiol. 2017 Jul 25;8:1402. doi: 10.3389/fmicb.2017.01402. PMID: 28790995; PMCID: PMC5524727.

-Dr. Richard Davis, PhD, D(ABMM), MLS(ASCP)CM is a clinical microbiologist and regional director of microbiology for Providence Health Care in Eastern Washington. A certified medical laboratory scientist, he received his PhD studying the tropical parasite Leishmania. He transitioned back to laboratory medicine (though he still loves parasites!), and completed a clinical microbiology fellowship at the University of Utah/ARUP Laboratories in Utah before accepting his current position. He is a 2020 ASCP 40 Under Forty Honoree.

Microbiology Case Study: A 30 Year Old with Fever Post Stem Cell Transplant

Case History

A thirty year old female with refractory acute leukemia was admitted to undergo allogeneic stem cell transplantation. Her initial admission had multiple infectious complications and chemotherapy-induced pancytopenia with profound absolute neutropenia. The patient was placed on prophylaxis/therapy including bacterial, viral, and fungal coverage. On hospital day 14, the patient was febrile to 38°C; vancomycin/piperacillin-tazobactam were added as empiric therapy due to concern for sepsis with fluctuation in mental status. CT Brain without contrast revealed a large intracranial hematoma with mass effect. Her mental status continued to decline and intubation was required for airway protection. An emergent decompressive hemicraniectomy was performed where necrotic brain tissue with hemorrhage/clot were found.

Due to ongoing fevers, empiric antimicrobial therapy was further broadened with meropenem, doxycycline, trimethoprim/sulfamethoxazole, and liposomal amphotericin B (L-AMB). Repeat cultures and imaging studies were ordered to evaluate for infection as a fever source. CT Angiography Chest (Image 1A-C) was performed and revealed an extensive non-enhancing area of ground glass opacity with peribronchovascular consolidation (“Reversed Halo” sign) in the right lung concerning for angioinvasive fungal infection. A tracheal aspirate was sent for bacterial and fungal culture. Subsequent bronchoscopy revealed extensive necrosis involving all visualized airways of the right tracheobronchial tree to the first subsegmental level (Image 1D). By contrast, the left lung appeared relatively normal and uninvolved. Bronchial washings of the right lung were also submitted for culture.

Image 1. Computed Tomography (CT) Angiography Chest images: Coronal (A), Cross-section (B) and Sagittal (C) sections reveal a large central ground glass opacity surrounded by a dense consolidation in the shape of a ring “Reversed Halo” sign. Bronchoscopy (D) revealed extensive necrosis and friable mucosa in the visible airways of the right lung.

Laboratory Identification

Respiratory specimens were sent to the microbiology laboratory for bacterial and fungal cultures. Hyphal elements were observed on the Gram stain of the submitted tracheal aspirate (Image 2A). Robust fungal growth was noted within 48 hours on Brain Heart Infusion and Inhibitor Mold agars from both the tracheal aspirate and bronchial wash specimens (Image 2B). A lactophenol cotton blue prep revealed broad hyphae with few septations, consistent with a member of the Mucorales (Image 3). Sporangiophores were noted to be long, dark and branched with round sporangia. Few rhizoids were observed and located between the sporangiophores. A definitive identification of Rhizomucor sp. was obtained through the use of matrix-assisted laser desorption ionization mass spectrometry (MALDI-TOF MS).

Image 2. A: Gram stain (400X) of direct sample from tracheal aspirate with broad, irregular, long, pauciseptate hyphae consistent with infection from a fungal species belonging to the order Mucorales. B: Fluffy “cotton candy” appearing light brown-grey colonies grew rapidly (<4 days) on brain heart infusion agar from both tracheal aspirate (pictured) and bronchial washing specimens.
Image 3. Lactophenol cotton blue stain obtained by tape preparation direct from fungal colony (A; 200X, B-D; 400X). Numerous branched dark brown non-apophysate sporangiophores with spherical columella were seen along with numerous broad, irregular pauciseptate hyphae with right angle branching (A, B). Rudimentary rhizoids developing off stolons (C, D) were rarely identified.


Members of the order Mucorales can be identified in the laboratory by their rapid, robust growth and cotton candy-like appearance on conventional fungal media excluding those containing cycloheximide.2 Microscopically, these molds exhibit pauciseptate, broad (9-15 μm wide) hyphae with sporangia. Some species elaborate root-like structures called rhizoids (e.g Rhizopus, Rhizomucor) while others lack them (e.g. Mucor).2 Rhizomucor sp. can be differentiated microscopically from other related members of the Mucorales by its branched, dark brown sporangiophores with absent apophysis, round columella and the presence of few, short, rudimentary rhizoids (Image 3A-D). In practice, the rhizoids can be challenging to identify microscopically.2 Some species can also grow at elevated temperatures (~38-58°C) which can be utilized as a tool for use in identification.2 Newer methods including the use of MALDI-TOF MS and DNA probes allow for rapid, accurate identification of these fungi, but are not routinely available in all laboratories.

Infections caused by Mucorales (Mucormycosis) usually involve immunocompromised patients with defects in cell-mediated immunity. Rapid and often fatal, these infections can prove extraordinarily difficult to manage. They are known to be angioinvasive and can widely disseminate. Debridement of involved tissues and higher-level antifungal agents (e.g. posaconazole, amphotericin B) are mainstays of therapy. The most recognized group of patients where these infections are identified are uncontrolled diabetics, frequently in diabetic ketoacidosis, often with nasal/orbital sinus involvement. Patients with leukemia/lymphoma who are undergoing stem cell transplantation are another group often affected. In addition to the nasal/orbital sinuses, the gastrointestinal tract, skin and lungs also serve as important sites for these infections, especially if mucositis occurs.1

Rhizomucor sp. are an occasional cause of mucormycosis, but have a predilection for leukemic patients such as in this case.2 Given the significant bronchoscopy findings and the intraoperative presence of necrotic brain tissue, there was substantial clinical concern for invasive pulmonary mucormycosis with possible central nervous system involvement. Isavuconazole was discontinued, and posaconazole and micafungin were added to her antifungal therapy (L-AMB). Granulocyte infusions were used in an attempt to increase her cell-mediated immune response to the mold. Cardiothoracic surgery evaluated the patient but the lesion was deemed unresectable. Due to the presence of epistaxis, the otolaryngology service evaluated the patient for invasive fungal sinusitis; however, nasal endoscopy did not reveal any nasal/sinus involvement. The patient never regained significant neurological function and continued to medically decline during the hospitalization. She was placed on comfort care where she died shortly afterwards.


  1. Love GL, Ribes JA. 2018. Color Atlas of Mycology, An Illustrated Field Guide Based on Proficiency Testing. College of American Pathologists (CAP), p. 244-274
  2. Walsh TJ, Hayden RT, Larone DH. 2018. Larone’s Medically Important Fungi, A Guide to Identification. ASM Press, p. 185-190

-John Markantonis, DO is the current Medical Microbiology fellow at UT Southwestern and will be completing his Clinical Pathology residency in 2022. He is also interested in Transfusion Medicine and parasitic diseases.

Kim Stewart BS, MT(ASCP)SM holds a bachelor’s degree from Texas Tech University and is medical technologist in the microbiology section at UT Southwestern Medical Center with 35 years’ experience.

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

Microbiology Case Study: 68 Year Old Female with Streptococcus anginosus in a Tracheostomy Wound

Case History

A 68 year old female was admitted to our hospital for emergent tracheostomy due to airway obstruction. Her symptoms began months ago with cough, dysphagia, and hoarseness of voice, difficulty breathing, slowly growing neck mass, and weight loss. Surgical biopsy from the neck mass confirmed the presence of squamous cell carcinoma of the hypopharynx “stage IVB”.

Chest CT scan indicated the possibility of metastatic disease to the lungs. Recently, she experienced a slow onset of paraparesis and blurring of vision in her left eye, which raised a concern about the disease metastasized to the brain. Brain MRI was performed and came back negative for metastasis to the brain. Additionally, she is at risk of refeeding syndrome due to muscle wasting, cachexia, dysphagia, 25% body weight loss, and significant failure to thrive. Radiation oncologists recommended concurrent radiation with chemotherapy, with a prescribed 70 Gray of radiation dose.

Physical examinations showed bilateral coarse lung sounds. Prior to her presentation, she had been treated with docusate, hydrocodone-acetaminophen, morphine, ondansetron, and labetalol. Her past medical history was notable for essential hypertension and an extensive history of smoking 1 PPD for 45 years. She was also started on weekly carboplatin and paclitaxel for five weeks.

The surgical biopsy from the tracheostomy lesions was sent to the microbiology laboratory for bacterial culture. The Gram stain of the culture showed gram positive cocci in chains. After 48 hours of incubation, the cultures grew a pure culture of α-hemolytic colonies (Figure 1). The organism was identified as Streptococcus anginosus by MALDI-TOF mass spectrometry (VITEK MS).

Image 1. Alpha-hemolytic colonies of Streptococcus anginosus growing on Sheep Blood Agar (SBA) plate after 48 hours of incubation.


Streptococcus anginosus group (SAG) “formerly named as streptococcus milleri” are members of the Viridians Streptococci group, which are known to cause endocarditis due to their ability to bind extracellular matrix proteins such as fibronectin, fibrinogen, and laminin, by facilitating bacterial adhesion to the heart valves.1

In general, SAG consists of three main strains: Streptococcus intermedius, Streptococcus constellatus, and Streptococcus anginosus. Streptococcus intermedius tends to be associated with central nervous system (CNS) infections, while S. anginosus is commonly found as commensals at the genitourinary and gastrointestinal systems.2 Since S. anginosus strains can be virulent due to its ability to survive in an acidic environment and cause systemic bacteremia, skin and soft tissue infections (SSTIs), osteomyelitis, and CNS infections, isolation of this organismfrom invasive sites should not be regarded as a contaminant. Besides, S. anginosus infection is occasionally associated with liver abscesses and colonic adenocarcinoma.3

Notably, the recovery of this organism from our patient’s tracheostomy biopsy wound indicates the likely association of S. anginosus and oral squamous cell carcinoma (OSCC). Sasaki M et al. demonstrated that the dental plaques in oral squamous cell carcinoma patients could act as a reservoir for SAG, which might cause significant DNA damage in oral mucosa, thus predisposing to accumulated mutations.4 Two other studies have also shown that SAG is recovered exclusively in oral squamous cell carcinoma patients compared to individuals without oropharyngeal cancer.5,6

SAG are usually susceptible to penicillin, ampicillin, or ceftriaxone, while sometimes they can be resistant to clindamycin.7 In our case, the patient received multiple doses of IV ampicillin-sulbactam and metronidazole in the emergency department. After S. anginosus had been identified from her tracheostomy wound, the patient was discharged on oral ampicillin-sulbactam, along with the carboplatin and paclitaxel treatment.


  1. Asam D, Spellerberg B. Molecular pathogenicity of Streptococcus anginosus. Mol Oral Microbiol. 2014;29(4):145-155. doi:10.1111/omi.12056
  2. Whiley RA, Beighton D, Winstanley TG, Fraser HY, Hardie JM. Streptococcus intermedius, Streptococcus constellatus, and Streptococcus anginosus (the Streptococcus milleri group): association with different body sites and clinical infections. J Clin Microbiol. 1992;30(1):243-244. doi:10.1128/JCM.30.1.243-244.1992.
  3. Masood U, Sharma A, Lowe D, Khan R, Manocha D. Colorectal Cancer Associated with Streptococcus anginosus Bacteremia and Liver Abscesses. Case Rep Gastroenterol. 2016;10(3):769-774. Published 2016 Dec 13. doi:10.1159/000452757
  4. Sasaki M, Yamaura C, Ohara-Nemoto Y, et al. Streptococcus anginosus infection in oral cancer and its infection route. Oral Dis. 2005;11(3):151-156. doi:10.1111/j.1601-0825.2005.01051.x
  5. Robayo DAG, Erira HAT, Jaimes FOG, Torres AM, Galindo AIC. Oropharyngeal Squamous Cell Carcinoma: Human Papilloma Virus Coinfection with Streptococcus anginosus. Braz Dent J. 2019;30(6):626-633. doi:10.1590/0103-6440201902805.
  6. Tateda M, Shiga K, Saijo S, Sone M, Hori T, Yokoyama J, Matsuura K, Takasaka T, Miyagi T. Streptococcus anginosus in head and neck squamous cell carcinoma: implication in carcinogenesis. Int J Mol Med. 2000 Dec;6(6):699-703. doi: 10.3892/ijmm.6.6.699. PMID: 11078831.
  7. Tracy M, Wanahita A, Shuhatovich Y, Goldsmith EA, Clarridge JE 3rd, Musher DM. Antibiotic susceptibilities of genetically characterized Streptococcus milleri group strains. Antimicrob Agents Chemother. 2001;45(5):1511-1514. doi:10.1128/AAC.45.5.1511-1514.2001.

-Ahmed Ismail Younes, MD., is a first year pathology (AP/CP) resident at the East Carolina University Brody School Medicine. He is interested in specializing in dermatopathology. He is also passionate about conducting translational research. In his spare time, Ahmed enjoys spending his time watching movies, baking homemade pizza, and playing soccer.

-Phyu M. Thwe, Ph.D., MLS (ASCP)CM is Technical Director/Technical Consultant at Vidant Medical Center Clinical Microbiology Laboratory. She completed a Clinical and Public Health Microbiology Fellowship through a CPEP-accredited program at the University of Texas Medical Branch (UTMB) in Galveston, Texas. She is interested in extrapulmonary tuberculosis pathophysiology and developing diagnostic algorithms.

Microbiology Case Study: Interesting Case of a Cavitary Lung Mass

Case History

A 50 year old male with a significant past medical history of poorly controlled type 2 diabetes mellitus, hypertension, hyperlipidemia, smoking tobacco abuse and obstructive sleep apnea was referred to our institution’s pulmonology clinic for cavitary lung mass. The lung mass was incidentally discovered on chest x-ray and has been clinically stable on serial imaging for over two years; however, a previous extensive laboratory workup including computed tomography (CT) guided biopsy was unrevealing to its etiology. The patient was noted to be largely asymptomatic at his initial office visit; repeat diagnostic workup was ordered. CT chest imaging revealed a 2.8 x 1.9 x 2.0 cm cavitary lung mass in the posterior left lower lobe that was unchanged compared to outside CT imaging from approximately 4 months prior.

Image 1. Cross section (left) and Sagittal (right) views from CT chest without contrast revealed a 2.8 cm transverse by 1.9 cm anteroposterior by 2.0 cm craniocaudal stable mass-like opacity in the left lower lobe superior segment broadly abutting the posterior pleura with a tiny internal focus of cavitation.

Given the chronicity of the lung mass, atypical infection (Nocardia, endemic fungi, mycobacterium) and primary pulmonary cancer were highest on the differential diagnosis. Blood tests including endemic fungal serologies, QuantiFERON-TB Gold, cryptococcal antigen, galactomannan and Fungitell (1-3)-B-d glucan assay were negative. Given the unrevealing non-invasive workup, a repeat CT guided biopsy was performed and core biopsy samples were sent for AFB, fungal and Nocardia cultures as well as for histopathological examination.

Histopathology revealed necrotizing granulomatous inflammation with empty spherules of Coccidioides suggestive of a remote infection of long duration (Images 2, 3). Additionally, no microorganisms were isolated from cultures. Based on these findings, an infectious disease (ID) consult was placed. The patient remained asymptomatic and revealed a long history of residing within areas of the Southwestern United States endemic to Coccidioides species (sp.) during his ID office visit. Repeat Coccidioides complement fixation was positive for IgG (Titer: 1:4) with negative IgM by immunodiffusion testing. Urine Coccidioides antigen tested by quantitative sandwich enzyme immunoassay was negative. These findings likely represent past history of coccidiomycosis and not active infection. Antifungal therapy was deferred due to the patient’s asymptomatic status. The patient was monitored with close clinical follow up and continued serial imaging.

Histopathology Images

Image 2. Hematoxylin and eosin stained sections of formalin fixed paraffin embedded (FFPE) tissue from core biopsy of cavitary lung mass. Necrotizing granulomatous inflammation at 40X (A) and 100X (B) with empty spherules of Coccidioides (C, D) at 600X.
Image 3. Special stains performed of formalin fixed paraffin embedded (FFPE) tissue from core biopsy of cavitary lung mass highlighting empty spherules. Grocott’s methenamine silver stain at 100X (A) and 400X (B). Periodic Acid Schiff for Fungus stain at 600X (C).


Coccidioides sp. are dimorphic fungi with a mycelial (saprophytic) phase in the environment and a spherule (parasitic) phase in its host.1 It is the cause of coccidiomycosis, also known as valley fever, desert fever or San Joaquin fever, which has a wide range of clinical presentations from subclinical manifestations (~60%) to an influenza-like illness followed by skin lesions to the most pathological form, disseminated disease.1 It can also cause the development of cavitary lung masses, as described in this case.1 It is endemic to the Southwestern region of the United States where it prefers dry, arid conditions.2 Infections normally occur by inhalation of infective arthroconidia, which have matured from mycelium, following disruption of soil.1 Once inside the host, lungs spherules containing endospores develop (Image 4).1 These spherules rupture releasing the endospores which can continue to develop into spherules to maintain a continuous parasitic cycle or can be exhaled into the environment to continue its saprophytic phase.1

Image 4. High magnification images of hematoxylin and eosin stained sections of formalin fixed paraffin embedded (FFPE) lung tissue revealing multiple spherules containing endospores (left) consistent with active Coccidioides infection and a giant ruptured spherule releasing endospores (right) that will continue to propagate Coccidioides infection.

Two morphologically indistinct species exist (C. immitis and C. posadasii) that can only be definitively identified by molecular methods.3 C. immitis is predominantly found in central and southern California while C. posadasii can be found in other non-Californian southwestern US states and extending into western Texas and down throughout Mexico and South America.3 When cultured, it grows rapidly at both 25°C and 37°C into woolly white colonies that develop alternating barrel-shaped arthroconidia that can be seen on tape prep with lactophenol blue.4


  1. Donovan FM, Shubitz L, Powell D, Orbach M, Frelinger J, Galgiani JM. 2019. Early Events in Coccidiomycosis. Clinical Microbiology Reviews, 33, e00112-19, DOI: 10.1128/CMR.00112-19
  2. Hernandez H, Erives VH, Martinez LR. 2019. Coccidioidomycosis: Epidemiology, Fungal Pathogenesis and Therapeutic Development. Current Tropical Medicine Reports, 6, 132-144,  DOI: 10.1007/s40475-019-00184-z
  3. Kirkland TN, Fierer J. 2018. Coccidioides immitis and posadasii; a review of their biology, genomics, pathogenesis, and host immunity, Virulence, 9:1, 1426-1435, DOI: 10.1080/21505594.2018.1509667
  4. Love GL, Ribes JA. 2018. Color Atlas of Mycology, An Illustrated Field Guide Based on Proficiency Testing. College of American Pathologists (CAP), p. 234-235

-John Markantonis is the current Medical Microbiology fellow at UT Southwestern and will be completing his Clinical Pathology residency in 2022. He is also interested in Transfusion Medicine and parasitic diseases.

-Dominick Cavuoti is a Professor at UT Southwestern and specializes in Infectious Diseases Pathology, Medical Microbiology 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.

What to Expect When You Don’t Know What You’re Expecting: COVID-19 and Flu Season in the Laboratory

Welcome to October 2020 and a flu season unlike any other. What can we expect? Well, it’s complicated. And if we aren’t sure what to expect, can we still be prepared? Yes (at least for some things)!

From the beginning of the COVID-19 pandemic and throughout the summer of 2020 clinicians and laboratorians have been anxiously wondering what effect global presence of respiratory virus SARS-CoV-2 would have on the 2020-2021 flu season. “Flu season,” the annual, relatively predictable period of increased cases and deaths due to Influenza A and B, occurs during colder, winter months. In the northern hemisphere this is September through March. We have extensive experience tracking the onset and genetic variability of the predominant influenza viruses. We manufacturer flu vaccines based on data of potentially likely influenza strains. Other viruses that cause respiratory symptoms follow similar seasonal patterns. These include common (non-SARS-CoV-2) human coronaviruses, and Respiratory Syncytial Virus (RSV). In short: this is a known, annual occurrence that we can usually prepare to some extent.

So what will that look like this year? During the historic 1918 pandemic influenza, deaths seen during the first winter of the outbreak paled in comparison to those seen the following winter. Even if that kind of terrible scenario doesn’t occur during this pandemic year, it is possible we are facing “perfect storm” of COVID-19 plus influenza resulting in overwhelmed hospitals and depleted testing supplies. [https://www.cidrap.umn.edu/news-perspective/2020/09/fears-perfect-storm-flu-season-nears]

We know that COVID-19 spreads well in enclosed spaces with prolonged person-to-person contact, regardless of climate and temperature, via respiratory secretions. Because of this, there has been widespread adoption of mask wearing, social distancing, and limitations on in-person gathering. Promisingly, these interventions to prevent the spread of COVID-19 seem to be contributing to historically low influenza rates in the Southern Hemisphere! [https://www.cdc.gov/mmwr/volumes/69/wr/mm6937a6.htm] But adoption of these mitigation strategies are not being universally or rigorously followed in all regions and communities. As temperatures drop, we could see more people conducting activity indoors – will this change transmission patterns? Will regions with ongoing COVID-19 outbreaks be more prone to influenza as well? If hospital capacity becomes strained, will criteria for ordering tests change?

During COVID-19 laboratories have responded heroically and rapidly to test kit shortages, supply chain issues, and staffing challenges. At this stage (October of 2020) many high-level decisions about SARS-CoV-2 testing, like test platform purchasing and validation or manufacturer test kit allocations, might already be set in stone. So is there anything that can be done to help labs and laboratory workers successfully make it through flu season?

Here are 3 suggestions:

1) Establish testing algorithms and clear sample workflows.

Each facility and laboratory will have their own platforms for testing COVID-19 and other respiratory pathogens. Depending on the service ordering the test, there can be both immediate and downstream consequences for when a test comes back positive, negative, or even when that test result is slower than expected!

An algorithm helps set institutional expectations for what tests are ordered under different scenarios. For example symptomatic patients presenting to a hospital with influenza-like illness (ILI), especially when they will be admitted, should likely have both SARS-CoV-2 and influenza tests ordered simultaneously. But asymptomatic patients being admitted for procedures may only require a SARS-CoV-2 test.

Let’s say your lab has both a SARS-CoV-2 PCR test and SARS-CoV-2 rapid antigen test. But due to risk a false negative, lab and clinical leaders are uncomfortable using only a rapid antigen test to conclusively rule out COVID-19 in patients being admitted to the hospital. Your algorithm could use specify the use of SARS-CoV-2 antigen testing in symptomatic patients to quickly “rule in” potential positives, where antigen-negative patients will also have a PCR test. Algorithm specifics come down to what your institutions stake holders (clinical AND laboratory) need and capacity are. The details of an algorithm will be dependent on your lab test platforms, your available test orders, and may need to be modified to accommodate restricted test allocations.

Along with clinical algorithms, clear workflow for specimens and test types can help laboratory workers get tests where they need to go within the lab. Not all SARS-CoV-2 tests have approval in the instructions for use for, say, nasal swabs. If nasal swab comes to the lab with orders for both influenza and SARS-CoV-2 tests, what is the procedure for informing the floor for an appropriate collection? Or say that your test platforms for different tests live in different areas of the lab. Your workflow may be to set up one test and do a pour off into an aliquot tube so tests can be run at the same time. Or you may have sufficient test collection materials to request a separate sample for each test.

Probably the most important part of developing or reviewing your existing algorithms and laboratory workflow is doing it in connection with others. The purpose is to streamline the entire process from clinical decision making to test performing and reporting and help everyone be on the same page.

2) Communicate to clinical staff frequently about your tests.

Because of the intense interest surrounding COVID-19 laboratory testing, it’s entirely possible that more people have had to learn about previously niche laboratory concepts like “sensitivity vs. specificity” and “PCR vs. antibody vs. antigen tests” than at any previously time in human history! However, it is also likely that many clinicians or administrators in your own institution may know more about a test platform they read about in the news than the COVID-19 test platform that their laboratory performs.

Even at this stage in the pandemic with perhaps more exposure (pun not intended!) then the laboratory has ever had, miscommunication and unclear expectations abound surrounding test performance or turnaround times.

Whenever possible, lab leaders who interact with clinicians and administrators should look for ways to educate on test platforms, testing capacity, and expected test performance (i.e. time to result, comparative sensitivity etc.). This could include asking for time to provide formal updates during monthly meetings, monitoring test statistics (e.g. a test “dashboard”), or just informal reminders about what tests the lab performs during phone calls.

3) Keep the lab staff off the phone.

A critical part of the job of the lab is to provide information and updates on when test results are available. But when the hospital floors or clinics are busiest with patients, often the lab is busiest performing those patients’ tests. A phone call about the status of a respiratory virus test can be undeniably helpful to that patient’s clinical care team! But a dozen such phone calls over the course of a lab worker’s shift, especially under normal lab conditions (e.g. no staff shortages or instrument issues) is a failure of communication and can be detrimental to both lab performance and lab worker wellbeing.

In addition to the need for regular education about testing mentioned above, to help protect your lab staff’s bench time here are some possible ways keep from being overwhelmed with phone calls:

  • In some institutions, passive reminders (for example about hand hygiene or upcoming events) cycle through computer screen savers or on television screens in clinical areas. You could see if a message like “Reminder from the lab: COVID-19 tests are completed in [length of time].” could be put on a rotation.
  • If there is no client service or switchboard for your lab, but people call the lab directly for updates, you could institute a message stop. This is where phone calls routed to the laboratory must listen to a reminder that (for example), “If you are calling for an update of a COVID-19 test, these tests cannot be completed faster than [length of time] after arriving in the lab.”

    While these messages can be undeniably annoying and disruptive for people calling the lab for other reasons (and become less effective over time) if phone calls get out of hand, this option could be considered.
  • A lab instrument going down can result in test backlogs and numerous phone calls to the lab. Some institutions centralize their information in the form of a duty officer (for example in the emergency department). This will be a person who can be informed of actionable information, like test delays due to instrument issues, and who will post and distribute that information to those affected.

There is a lot we don’t know about what’s to come in the COVID-19 pandemic. While we can’t predict the ways the lab may be challenged with the next unforeseen disruption, or even what our flu season testing needs may look like, hopefully we can prepare now to continue to support our patients by helping and supporting our labs.

-Dr. Richard Davis, PhD, D(ABMM), MLS(ASCP)CM is a clinical microbiologist and regional director of microbiology for Providence Health Care in Eastern Washington. A certified medical laboratory scientist, he received his PhD studying the tropical parasite Leishmania. He transitioned back to laboratory medicine (though he still loves parasites!), and completed a clinical microbiology fellowship at the University of Utah/ARUP Laboratories in Utah before accepting his current position. He is a 2020 ASCP 40 Under Forty Honoree.

Microbiology Case Study: A Middle-Aged Woman with Forearm Pain

Case History

A middle-aged female was evaluated for left forearm pain and erythema following a cat bite one-day prior, and was prescribed trimethoprim-sulfamethoxazole for management in the outpatient setting.  She subsequently presented for follow-up where she was noted to have a 3 x 4 cm raised, red, indurated lesion of left arm without any discharge (Image 1).  MRI demonstrated a 6.5 x 2.3 x 2.3 cm abscess within the distal ulnar soft tissues with surrounding cellulitis.  As her pain and erythema were progressively worsening, she was admitted for surgical management.   

Upon admission, a bedside incision found purulent drainage which grew mixed anaerobic gram negative rods.  Blood cultures collected at this same time were negative and remained so for the duration of her hospital course.  Empiric antibiotic therapy was initiated with piperacillin-tazobactam, and the patient underwent formal surgical incision and drainage.  Intraoperative findings were notable for abscess, diffuse and severe tendinopathy, and a thick inflammatory rind surrounding the associated neurovascular bundle.  Intraoperative cultures were obtained and sent to the microbiology laboratory.  The patient’s postoperative course was uneventful, and she was discharged with plans to complete a two week course of amoxicillin-clavulanate.  Follow-up clinic visits demonstrated successful recovery, with a well-healed incision and normal grip strength and range of motion.

Laboratory Identification

Bacterial culture of abscess material collected intraoperatively grew smooth, mucoid colonies on chocolate and blood agars with less than 24 hours of incubation at 35°C in CO2 (Image 2, bacterial isolate). Growth was notably absent on MacConkey agar. Gram stain of the colony revealed tiny, gram negative coccobacilli (Image 2). Biochemical testing determined this organism to be indole, oxidase, and catalase positive.  The organism was definitively identified as Pasteurella multocida by MALDI-TOF MS.

Image 1. Arm lesion prior to incision and drainage.
Image 2. P. multocida growth on Blood and Chocolate agars. Gram stain from a colony revealed small, gram negative coccobacilli (far right).


Members of the genus Pasteurella are small, Gram-negative coccobacilli which are able to readily grow on Sheep’s blood agar and chocolate agar, but will typically not grow on MacConkey media.  Infection with these organisms is usually considered to be a zoonosis, with both wild and domestic animals serving as reservoirs.  In animal hosts they can be part of the endogenous flora or pathogens.  P. multocida is the most common member of the genus associated with human infections, which has now been divided into multiple taxonomic subspecies through the use of more discriminatory molecular methods.  Biochemically, P. multocida is positive for catalase, oxidase, indole, and nitrate reduction.

Animal bite wounds are often polymicrobial and contain mixtures of both aerobic and anaerobic organisms.  These organisms can be reflective of the oral flora of the biting animal, or of endogenous skin flora of the bite victim.1  While 80-90% of bites per year can be attributed to dogs, an estimated 400,000 cat bites (5-10% of the total) occur in the United States annually.2  While dog bites often manifest as localized crush injuries with contusions and/or lacerations, a majority of such wounds are accessible to irrigation and cleaning which leads to a relatively low infection rate (5-10%).  By contrast, cat bites are often deep, localized puncture wounds which provide excellent environments for the growth of both aerobic and anaerobic bacteria.  While feline bite wounds may appear less severe after cursory inspection, these wounds can be considerably more difficult to clean, resulting in overall infection rates up to 50%.3 

Management of bite wounds includes cleansing, irrigation and debridement.  Importantly, antimicrobial therapy should include coverage for both aerobic and anaerobic bacteria.4  In this case, amoxicillin-clavulanate was utilized with good results, and provides coverage for the most common oral aerobes and anaerobes encountered in animal bite wounds.  Amoxicillin-clavulanate also has activity against beta-lactamase producing bacteria such as Prevotella sp. and Porphyromonas sp. which are oral anaerobes of dogs, cats, and humans.  The use of macrolides should be avoided due to variable activity against Pasteurella multocida.4  As in this case, bite wounds most frequently are encountered on the upper extremities, and Pasteurella sp. is one of the most common isolates recovered from bites from both cats and dogs (50% of dog bites, and 75% of cat bites).2


1. Abrahamian FM, Goldstein EJC. 2011. Microbiology of Animal Bite Wound Infections. Clinical Microbiology Reviews 24:231.

2. Bula-Rudas FJ, Olcott JL. 2018. Human and Animal Bites. Pediatrics in Review 39:490.

3. Kannikeswaran N, Kamat D. 2008. Mammalian Bites. Clinical Pediatrics 48:145-148.

4. 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. Clinical Infectious Diseases 59:e10-e52.

 -Francesca Lee, MD, is an associate professor in the Departments of Pathology and Internal Medicine (Infectious Diseases) at UT Southwestern Medical Center.

-Huy Dao, MLS(ASCP)CM graduated from the University of Minnesota and has worked for eight years as medical technologist for eight years.  He is interested in clinical mycology and bacteriology.

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

Microbiology Case Study: 83 Year Old Female with a Perisplenic Abscess

Case History

An 83 year old female with a past medical history of breast cancer, multiple strokes, dysphagia, hypertension and gastroesophageal reflux disease (GERD) presented to an outside hospital with altered mental status, metabolic encephalopathy, decreased appetite, acute kidney injury, and E. coli/Proteus urinary tract infection (UTI). There, she was diagnosed with a perforated gastric ulcer, which was repaired, with a gastrostomy (G) tube in place. The patient later developed a fever and an aspiration pneumonia, which was treated with ampicillin/sulbactam. A follow up imaging revealed a new gastric perforation along the fundus of the stomach with perisplenic fluid collection, along with a pleural effusion with possible communication with the fluid collection in the stomach. Due to her complex medical history, an additional intervention was not pursued and the family agreed to comfort measures, and the patient was discharged home.

The family presented to our emergency department the same day of discharge, as the patient had not been eating and the family needed assistance in using the G tube. In-house computed tomography (CT) of the abdomen/pelvis again showed an organizing collection near the spleen (Image 1). A medium-sized left pleural effusion with left lower lobe collapse due to the communication with the perforation was observed on CT. The patient received IV fluids and a dose of vancomycin and zosyn in the ED. A vascular and interventional radiology (VIR) consult was recommended for potential drainage of the perisplenic abscess and left pleural effusion.

Image 1. CT of the abdomen demonstrating an organizing collection (black circle, approximately 7 cm x 3 cm) posterior/superior to the spleen near the site of the prior gastric perforation concerning for an abscess.
Image 2. Small gram negative rods on a Gram stain of Burkholderia cenocepacia from a subculture.
Image 3. Culture morphology of Burkholderia cenocepacia on a blood agar plate after overnight incubation: smooth colonies are present (upon prolonged incubation, the colonies turned yellowish color – not shown in image).

VIR drained the perisplenic abscess, which was sent to the microbiology lab for aerobic & anaerobic cultures. The Gram stain revealed many white blood cells (WBC’s) and rare yeast. The culture grew 3+ Burkholderia cepacia complex (Burkholderia cenocepacia) and 3+ Candida glabrata. The Gram stain and colony of the subculture of B. cepacia on a blood agar plate are shown in Images 2 and 3. C. glabrata was also isolated from the urine culture. Susceptibility testing of B. cepacia showed that it was sensitive to both meropenem and trimethoprim-sulfamethoxazole. Vancomycin and zosyn were stopped and the patient was switched to IV trimethoprim sulfamethoxazole for B. cepacia and IV micafungin for C. glabrata.

Due to the recurrent perforation, the G tube could not be used; a jejunostomy (J) tube was placed instead. Feeds were successfully started with the J tube. Trimethoprim-sulfamethoxazole was also given via the J-tube. A follow-up endoscopy showed a normal esophagus, the known perforation in the gastric fundus, and erythematous duodenopathy at the level of the duodenal bulb, the remainder of the duodenum was normal. The patient’s clinical status improved and she was discharged home with the support of home health services.


We present an uncommon presentation of Burkholderia cenocepacia, a member of B. cepacia complex, in perisplenic abscess fluid. B. cepacia complex consists of at least 17 closely related species. They are rod-shaped, aerobic, motile Gram-negative bacteria. B. cepacia complex has been well characterized as opportunistic pathogens, particularly in patients with cystic fibrosis and chronic granulomatous disease (CGD). However, they can also infect immunocompetent patients and have been reported to cause endocarditis (specifically in IV drug abusers), pneumonitis, UTIs, osteomyelitis, dermatitis, and other wound infections. In the United States, B. multivorans and B. cenocepacia together account for approximately 80% of B. cepacia complex infections. Burkholderia have been isolated on contaminated hospital equipment and even disinfectants. They present a large problem in nosocomial infections due largely to their ability to survive in aqueous environments.1 They are soil-dwelling bacteria commonly found on plant roots. They are of environmental interest secondary to their antifungal and antinematodal properties as well as the ability to degrade many toxic compounds in agriculture (pesticides, herbicides, preservatives).2 Clinically important Burkholderia species outside of the B. cepacia complex include B. pseudomallei, the causative agent of melioidosis, and B. mallei, the causative agent of glanders.1

Rare case reports have previously documented B. cepacia isolated from splenic abscesses/infections. Most B. cepacia splenic infections occurred secondary to pneumonia or multi-organ involvements in CGD patients. 3, 4, 5 However, one report indicated the B. cepacia-mediated multiple splenic abscesses, in the setting of malignancy and diabetes. 6 While the splenic abscesses in the context of meliodosis, either due to B. pesudomallei or B. mallei infection, have been reported, 7 B. cepacia-mediated splenic infections are rarely encountered.

B. cepcacia complex has intrinsic resistance toseveral antibiotics including penicillins, amoxicillin-clavulanate, ertapenem, polymixin B, Colistin, and fosfomycin.8 B. cepacia complex possesses an inducible β-lactamase, encoded by the gene penA, which can hydrolyze penicillin and use it as a source of carbon. In one study involving 40 bloodstream isolates of B. cepacia in patients without cystic fibrosis, 93% of the isolates were susceptible to ceftazadime and 95% of isolates were susceptible to trimethoprim-sulfamethoxazole.9 Following discussion with our infectious disease colleagues, we believe that the B. cepacia isolate from our patient was likely a nosocomial infection from possible contamination of her G-tube in combination with the gastric perforation.


  1. Jorgensen, J. H., Pfaller, M. A., &amp; Carroll, K. C. (2015). Manual of clinical microbiology. Washington, DC, DC: ASM Press.
  2. Kenyon College Department of Biology. (2011, April 22). Burkholderia cepacia. Retrieved September 21, 2020, from https://microbewiki.kenyon.edu/index.php/MicrobeWiki
  3. Clegg HW, Ephros M, Newburger PE. Pseudomonas cepacia pneumonia in chronic granulomatous disease. Pediatr Infect Dis. 1986 Jan-Feb;5(1):111. PMID: 3945563.
  4. Sirinavin, Sayomporn MD*; Techasaensiri, Chonnamet MD*; Pakakasama, Samart MD*; Vorachit, Malai DSc; Pornkul, Rattanaporn MD; Wacharasin, Rames MD Hemophagocytic Syndrome and Burkholderia cepacia Splenic Microabscesses in a Child With Chronic Granulomatous Disease, The Pediatric Infectious Disease Journal: September 2004 – Volume 23 – Issue 9 – p 882-884 doi: 10.1097/01.inf.0000137565.23501.03
  5. Bottone EJ, Douglas SD, Rausen AR, Keusch GT. Association of Pseudomonas cepacia with chronic granulomatous disease. J Clin Microbiol. 1975 May;1(5):425-8. doi: 10.1128/JCM.1.5.425-428.1975. PMID: 1176612; PMCID: PMC275137.
  6. Jayawardena, M. N., Chandrasiri, N. S., Wijekoon, S., Madanayake, P., Corea, E., Ranasinghe, D. D., & Lamahewage, N. D. (2017). Burkholderia cepacia; an unusual cause of multiple splenic abscesses : A case report. Sri Lankan Journal of Infectious Diseases, 7(2), 123. doi:10.4038/sljid.v7i2.8146
  7. Chen, H., Hu, Z., Fang, Y., Lu, X., Li, L., Li, Y, Mao, X, Qian, L. (2018). Splenic abscess caused by Burkholderia pseudomallei. Medicine, 97(26). doi:10.1097/md.0000000000011208
  8. Patel, J. B., Weinstein, M. P., Eliopoulos, G.M., Jenkins, S.G., Lewis, J.S., Limbago, B., Mathers, A., Mazzulli, T., Patel, R., Richter, S.S., Satlin, M., Swenson, J.M., Traczewski, M.M., Turnidge, J.D. & Zimmer, B.L. (2017). Performance standards for antimicrobial susceptibility testing. Wayne, PA: Clinical and Laboratory Standards Institute.
  9. Bressler A.M., Kaye K.S., LiPuma, J.J., Alexander, B.D., Moore, C.M., Reller, L.B. & Woods, C.W. Risk factors for Burkholderia cepacia complex bacteremia among intensive care unit patients without cystic fibrosis: A case-control study. Infect Control Hosp Epidemiol 2007; 28(8):951-8 doi : http://dx.doi.org/10.1086/519177

-J. Stephen Stalls, MD is a PGY-II pathology resident at the East Carolina University Department of Pathology and Laboratory Medicine. He plans to pursue hematopathology and molecular pathology fellowships, but also greatly enjoys his time in the microbiology lab. Outside of work, he enjoys playing the drums and going to concerts.

-Phyu Thwe, PhD, MLS(ASCP)CM is Technical Consultant/Technical Director of Clinical Microbiology Laboratory at Vidant Medical Center, Greenville, NC.