microbiology – Page 23

Microbiology Case Study: A 59 Year Old Female with Fevers, Weakness, and Altered Mental Status

Case History

A 59 year old African American female presented to the emergency department with fevers, weakness, fatigue and altered mental status. Her past medical history was significant for hypertension, diabetes mellitus (type 2) with end stage renal disease and a recent cerebrovascular accident the month prior. Her surgical history included a mitral valve repair surgery three years ago and a renal transplant two years ago. Current medications included prednisone, mycophenolate and tacrolimus immunosuppressive agents. Physical examination was unremarkable except for a temperature of 101°F and she was oriented to person, place and time. Pertinent labs included a WBC count of 13.2 TH/cm², microcytic anemia, and a creatinine of 1.51 mg/dL. Due to previous cardiac surgery, a transesophageal echocardiograph (TEE) was performed and showed a large vegetation (1.6 x 1.5 cm) on the mitral valve. A diagnosis of endocarditis was made and the patient was started on broad-spectrum antibiotics & taken to surgery for a mitral valve replacement. Multiple blood cultures were negative to this point. Portions of the mitral valve were submitted to surgical pathology and the microbiology laboratory for bacterial, fungal and AFB cultures.

Laboratory identification

Surgical pathology received an aggregate of tan-yellow, fibrous tissue fragments (3.1 x 1.5 x 1.1 cm). Histologic assessment showed a confluent mass containing abundant narrow, septate hyphae consistent with a fungal infection (Image 1). No definitive pigment was identified. Grocott’s methenamine silver (GMS) stain also highlighted the narrow hyphae with numerous septations (Image 2). In the microbiology laboratory, a darkly pigmented mold grew after 5 days of incubation on Sabouraud dextrose agar (Image 3). Lactophenol cotton blue prep showed pigmented, curved conidia with 2-3 transverse septations consistent with Curvularia spp (Image 4). All blood cultures were finalized as no growth after 5 days. Fungitell was found to be greater than 500 pg/ml and Aspergillus galactomannan was negative (<0.5).

Image 1. Sections of mitral valve tissue showed a confluent mass of abundant hyphal elements (H&E, 4x).

Image 2. Special stains of the fungal mass highlighted narrow hyphae with numerous septations and acute angle branching (GMS, 4x).

Image 3. A darkly pigmented mold grew of Sabouraud dextrose agar after 5 days of incubation at 25°C.

Image 4. Many pigmented, curved conidia with multiple transverse septations were seen (lactophenol cotton blue prep, low power).

Discussion

Curvularia spp. belong to a heterogeneous group of dematiaceous or black molds. The presence of pigment in this category of molds is due to melanin in the hyphae. Dematiaceous molds are ubiquitous in nature and can occasionally cause human infections. These molds have a characteristic dark appearance on fungal media that is often dark gray, brown or black in color. In addition, when the reverse of the plate or slant is examined, the under surface is also pigmented. Based on their growth rate, the dematiaceous fungi are divided into the fast growers, such as Curvularia, Bipolaris and Alternaria spp., which are mature in 5-7 days. The second group is slow growers that take between 7-25 days to fully mature. Examples of slow growers include Phialophora, Exophila/Wangiella, Cladosporium, Fonsecaea and Rhinocladiella spp.

Most commonly, dematiaceous molds infections usually present as phaeophyphomycosis, chromoblastomycosis or mycetomas. These three entities are cutaneous or subcutaneous mycoses that are obtained by traumatic implantation but vary from one another based on clinical features and histologic features of the mold in tissues. They are most frequently cause infection in male agricultural workers in rural areas of tropical or subtropical climates. These infections are indolent in nature but can lead to significant morbidity over time, as they are difficult to treat effectively.

In addition to the above superficial infections, Curvularia spp. has also be known to cause keratitis, sinusitis and wound infections. In immunosuppressed individuals, disseminated infections with spread to the lungs and brain have been documented. Endocarditis due to Curvularia spp. is quite rare with very few cases previously reported in the literature. On those documented, Curvularia spp. infections tend to have a predilection for prosthetic heart valves or occur after cardiac surgery. Diagnosis of infective endocarditis is difficult as symptoms are indolent and blood cultures do not have a high yield. Therefore, culture of the vegetation may be the only way to make a diagnosis.

In the microbiology laboratory, Curvularia spp. will grow on routine fungal media as a darkly pigmented mold in a relatively short time. On lactophenol cotton blue prep, Curvularia spp. produce large conidia that usually contain 4 cells that are divided by transverse septations. The conidia take on a curved appearance due to swelling of the subterminal cell, which is often the largest and most deeply pigmented. If identification is necessary beyond the genus level, panfungal PCR assays followed by sequencing of ribosomal genes may be useful in providing a species level diagnosis from fresh or paraffin embedded tissue.

For localized infections, surgical treatment alone may be adequate in some cases. In infections that are extensive or if there is dissemination, treatment with newer triazoles, such as posaconazole or voriconazole, have shown a broad spectrum of activity against dematiceous molds. Amphotericin B is also another effective option. While susceptibility testing can be performed on clinically significant Curvularia spp. infections, interpretative breakpoints have not been defined and clinical correlation is lacking.

In the case of our patient, she remained on a ventilator following surgery and with the identification of mold on histology, she was started on micafungin. She was switched to amphotericin B after the mold was classified as Curvularia spp. Her condition did not improve despite therapy and she died 3 weeks after surgery.

-Azniv Azar, MD, is a fourth year anatomical and clinical pathology resident at the University of Mississippi Medical Center.

Stempak

-Lisa Stempak, MD, is an Assistant Professor of Pathology at the University of Mississippi Medical Center in Jackson, MS. She is certified by the American Board of Pathology in Anatomic and Clinical Pathology as well as Medical Microbiology. She is the director of the Microbiology and Serology Laboratories. Her interests include infectious disease histology, process and quality improvement, and resident education.

When Rapid Blood Culture Identification Results Don’t Correlate, Part 2: Contamination

All laboratories are prone to contamination events. Blood products, analyzers, reagents, media, etc. all have the potential to be contaminated. If you are a molecular microbiologist, then you have to worry about not only bacterial, but also nucleic acid contamination.

The Issue

The topic of my blog last month focused on discrepant results between blood culture and PCR. Traditional blood culture workflow involves correlating the Gram stain result to what grows in culture. Nowadays, many laboratories are also performing PCR on positive blood cultures. Because we know PCR is more sensitive, it may be easy for some to justify discrepancies. Let’s image that gram positive cocci in clusters were observed in the Gram stain, the PCR detected Staphylococcus and Enterococcus DNA, but only S. aureus grew in the culture. Where did the Enterococcus come from and where did it go? It was not observed in the Gram stain and it didn’t grow in cultures, so was it “real”? Possibly. It could be a contaminant or it could be real, just present in low numbers. It’s difficult to say without having to invest more effort.

When this type of situation occurs in my laboratory, three things happen. First, we review the data. For example, if the Gram stain is discrepant, then we review the Gram stain or perform an acridine orange stain (in the case of positive PCR, but negative culture). If it’s the PCR, then we would make sure that a result entry error did not occur, etc. Second, we add the comment, “clinical correlation needed”. We have found little value in going back to the blood culture bottle and trying to recover the missing organism because in most cases when we look hard enough, using selective agar and other strategies, we do find the organism from the PCR results buried among overgrowth. Therefore, our approach is to let the clinician know that they must use other clinical data to aid in their diagnosis. Third, we document all discrepant blood culture PCR results; which includes an automatic notification to the doctoral director.

Next, let’s imagine that two more blood cultures (from different patients) become positive all within a relatively short period of time from the first discrepant result noted above. gram negative bacilli are observed in one culture and the other displays gram positive bacilli. PCR detects Enterococcus DNA in both cases. What are the odds of that happening? Not good. Something strange is going on!

The Solution

A contamination investigation needs to immediately occur. The two likely sources of contamination are 1) the PCR assay or 2) the blood culture bottles. To determine whether the issue is due to amplicon or target contamination of the PCR assay, we need to identify which instruments reported the Enterococcus. Was it a single instrument or were different instruments involved? Our laboratory performs routine “swipe” tests of the environment as part of our quality control, which allows us to monitor contamination. Swipe tests may also be performed 1) after a known contamination event (i.e., spill due to cracked or leaky product) to ensure that decontamination was properly carried out, 2) to investigate increased positivity rates, or 3) follow up on unusual results, such as the scenario outlined above.

PCR may be performed on a random sampling of uninoculated bottles to determine whether the issue is due to contamination of the blood culture media. If the contamination is high density, this may be useful; however if it is low density, then all bottles you test may still be negative. If the contamination is due to bacterial DNA, then Gram stain or culture will not be useful, hence the need for PCR. It is important to note that the presence non-viable organisms and/or nucleic acids (at levels that can be detected by PCR) is a known limitation noted in the package insert of some blood culture media and PCR manufacturers. If contamination is suspected, then immediately file a report with the manufacturer. Be sure to document lot numbers and expiration dates so that they may alert other customers.
The Conclusion

Human error contributes to the majority of discordant laboratory results. However, errors in interpretation and result entry/clerical errors are only part of the problem. Contamination events only complicate matters. If the test volume is significant, then the number of discordant results should be quickly realized, especially if there truly is a contamination issue. It is important to have a process in place to help reconcile contamination events as quickly as possible as they have the potential to majorly impact operations and patient care.

References

https://labmedicineblog.com/2018/02/20/when-rapid-blood-culture-identification-results-dont-correlate-part-1-clinical-correlation-needed/

Martinez Headshot-small 2017

-Raquel Martinez, PhD, D(ABMM), was named an ASCP 40 Under Forty TOP FIVE honoree for 2017. She is one of two System Directors of Clinical and Molecular Microbiology at Geisinger Health System in Danville, Pennsylvania. Her research interests focus on infectious disease diagnostics, specifically rapid molecular technologies for the detection of bloodstream and respiratory virus infections, and antimicrobial resistance, with the overall goal to improve patient outcomes.

Hematology Case Study: The Race to Save a 48 Year Old Man from a Rare Disease

A 48-year-old Caucasian male presented to a Baltimore Emergency Room complaining of fever, chills, and aches. He stated he had not been feeling well for the past week. His symptoms had progressed rapidly over the last 3 days to include night sweats, nausea and excessive somnolence. History taken in the ER revealed the patient had returned 10 days prior from a Safari in Botswana and Zambia. The patient was admitted to the ICU, in shock, with a BP of 75/50. Even though the patient had taken anti-malarial medication, the doctors suspected malaria. Blood was sent to the lab for a blood parasite exam and treatment for malaria was started while the doctors waited for the confirmation.

In the Hematology laboratory, technologists perform microscopy of thick and thin blood smears to look for malarial parasites. The thin smear is a typical Wright Giemsa stained wedge smear, and the thick smears are prepared and stained so that the red blood cells are lysed, and the sample is concentrated, making examination easier. Thorough, careful examination of the thick smear is aimed to identify whether a particular parasite is present, but they require a long drying period and take several hours to prepare and read. Thin smears can detect the parasites but also permit identification of particular species of malaria. While the thick smears were drying the technologist examined the thin smear.

The technologist who examined this patient’s thin smears saw parasites (image 1) under her microscope. She consulted with a supervisor and pathologist to confirm, and the patient’s doctor was notified that the patient did not have malaria, but instead, had Trypanosoma! This was an exciting find in the laboratory, as there have been only 40 cases seen in the US in the past 50 years.

tryp1 — Image 1. This slide shows the parasite, in dark blue. The parasite causes
African trypanosomiasis, also known as sleeping sickness
(Courtesy of Greater Baltimore Medical Center).

The race for diagnosis and treatment did not stop there, as there are 2 types of African trypanosomiasis, or African sleeping sickness, and effective and appropriate treatment must be started in a timely fashion. Both types look identical on a blood smear and both are considered universally fatal, if not treated. West African trypanosomiasis and East African trypanosomiasis are caused by the tsetse fly, which only lives in rural Africa. The patient stated he did remember being bitten by tsetse flies, and because there had been such a short span of time between being bitten and the onset of symptoms, doctors concluded that the patient had the rarer and fast-acting East African trypanosomiasis, which can kill within months.

Epidemiologists at CDC were contacted, who then consulted other infectious disease specialists at CDC. There are 2 treatments depending the stage of the disease. Surinam is the first line of defense, but melarsoprol, which is arsenic-like and very toxic, must be used if the parasites have reached the central nervous system. Because of the urgent need to start treatment, emergency shipments of both drugs were flown to Baltimore. The patient was started on Surinam to reduce the number of parasites in his blood to a level low enough to allow a spinal tap to be performed. After confirming that the CSF showed no signs of the parasite, treatment with surinam was continued and the patient was discharged a week later and has made a full recovery.

Because of the excellent work done by the medical technologists who made the first discovery, the speed with which the critical calls were made, the actions of the doctors involved, and the cooperation of the CDC, this patient received his first dose of Surinam a little over 24 hours after his blood was sent to the lab. This case shows the importance of a thorough medical and travel history in differential diagnosis. It also illustrates the importance of the competency evaluations and surveys in which all laboratory professionals are required to participate. None of the technologists, doctors or scientists involved had ever actually seen a case of African Trypanosomiasis, they had only read about it in books and seen it on competency assessments.

This case is based on an actual case from 2016. My coworker, Gail Wilson, was the technologist who first saw the Trypanosoma on the slides. Many thanks to Gail for her keen eye and attention to detail!

tryp2 — Image 2: L&R: *Trypanosoma brucei* in thin blood smears stained with Giemsa. Center: A close up of a tsetse fly. Credit: DPDx

References

Jon E. Rosenblatt Barth Reller Melvin P. Weinstein.pages 1103-1108, Laboratory Diagnosis of Infections Due to Blood and Tissue Parasites Clinical Infectious Diseases, Volume 49, Issue 7, 1 October 2009; retrieved March 2018 from https://academic.oup.com/cid/article/49/7/1103/316703

Ivo Elliott, Trupti Patel, Jagrit Shah, and Pradhib Venkatesan. West-African trypanosomiasis in a returned traveller from Ghana: an unusual cause of progressive neurological decline BMJ Case Rep. 2014; 2014: bcr2014204451. Published online 2014 Aug 14.doi: 1136/bcr-2014-204451; retrieved March 2018 from https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4139564/

Lena H. Sun. Medical Detectives raced to save a man from a rare, ‘universally lethal’ disease; retrieved March 2018 from https://www.washingtonpost.com/news/to-your-health/wp/2016/12/22/medical-detectives-raced-to-save-a-man-from-a-rare-universally-lethal-disease/?utm_term=.16d7b136bc47

Parasites – African Trypanosomiasis (also known as Sleeping Sickness). Retrieved March 2018 from https://www.cdc.gov/parasites/sleepingsickness/

DPDx- Laboratory Identification of parasites of Public Health Concern; retrieved March 2018 from https://www.cdc.gov/dpdx/

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-Becky Socha, MS, MLS(ASCP)^CMBB^CMgraduated from Merrimack College in N. Andover, Massachusetts with a BS in Medical Technology and completed her MS in Clinical Laboratory Sciences at the University of Massachusetts, Lowell. She has worked as a Medical Technologist for over 30 years. She’s worked in all areas of the clinical laboratory, but has a special interest in Hematology and Blood Banking. When she’s not busy being a mad scientist, she can be found outside riding her bicycle.

Microbiology Case Study: A 52 Year Old Man with End Stage Renal Disease

Case History

A 52-year-old man with multiple medical issues including a history of end stage renal disease on hemodialysis, chronic pancreatitis status post distal pancreatectomy, intravenous drug use through dialysis catheter, and multiple types of bacteremia presented with chills and abdominal pain. Labs on admission included a white blood cell count of 28.64 k/cmm, hemoglobin 8.8 g/dL, and platelets 581 K. He was diagnosed with a pancreatitis flare and admitted for pain management, with further labs drawn. After one day, he felt much better and was discharged with a pending blood culture to follow up on. At 61 hours, one bottle flagged positive with yeast seen on gram stain.

Laboratory findings

cryptlaur1 — Image 1: potato flake agar with creamy tan-white colonies.

Image 2: calcofluor white fluorescent stain showing round yeast forms.

The organism was identified as Cryptococcus laurentii via MALDI-ToF MS. A follow-up fungal culture was negative, however, repeat blood culture grew Stenotrophomonas maltophilia. His tunneled catheter was removed, and two days later the patient required urgent interventional radiology access for dialysis. He completed a two-week course of ceftazidime and was discharged.

Discussion

Cryptococcus laurentii is a very rare fungal pathogen. It is a psychrophilic organism, growing ideally at 15 °C, and is the most common yeast found in tundra.¹ Major reservoirs include soil, food, and pigeon excrement.² C. laurentii usually causes infection in immunocompromised hosts, although rare incidents of infection in immunocompetent patients have been reported. Reported manifestations have included fungemia, meningitis, peritonitis, pneumonia, pyelonephritis, keratitis, and skin infection.³

Cryptococcus laurentii is a urease-positive organism. Gram stain shows large budding yeasts without hyphae. The yeast grows on routine agar as whitish-yellow creamy colonies and on birdseed agar as whitish or greenish colonies. Staining with calcofluor highlights encapsulated yeast forms. Molecular diagnosis can be accomplished by ribosomal RNA sequencing of the internal transcribed spacer and D1/D2 regions. Treatment in most cases has been with fluconazole, although in one case of peritoneal dialysis catheter-related peritonitis, voriconazole was used due to low fluconazole susceptibility.⁴

References

Molina-Leyva A, Ruiz-Carrascosa JC, Leyva-Garcia A, Husein-Elahmed H. Cutaneous Cryptococcus laurentii infection in an immunocompetent child. International Journal of Infectious Diseases. 2013;17(12). doi:10.1016/j.ijid.2013.04.017.
Johnson, L. B., Bradley, S. F. and Kauffman, C. A. Fungaemia due to Cryptococcus laurentii and a review of non-neoformans cryptococcaemia. Mycoses. 1998;41: 277–280. doi:10.1111/j.1439-0507.1998.tb00338.x
Furman-Kuklińska K, Naumnik B, Myśliwiec M. Fungaemia due to Cryptococcus laurentii as a complication of immunosuppressive therapy – a case report. Advances in Medical Sciences. 2009;54(1). doi:10.2478/v10039-009-0014-7.
Asano M, Mizutani M, Nagahara Y, et al. Successful Treatment of Cryptococcus laurentii Peritonitis in a Patient on Peritoneal Dialysis. Internal Medicine. 2015;54(8):941-944. doi:10.2169/internalmedicine.54.3586.

-Prajesh Adhikari, MD is a 3^rd year anatomic and clinical pathology resident at the University of Vermont Medical Center.

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

Microbiology Case Study: A 21 Year Old Female with a Sore Throat

A 21 year-old female presented to the emergency department with a sore throat. Her symptoms started two weeks prior to presentation. She was seen at student health last week and given Tamiflu, but her sore throat has grown progressively worse. She now has pain with swallowing and cannot swallow liquids. Upon examination the patient has no rash, no fever, and is not in respiratory distress. She does have left tonsillar fullness causing her uvula to be slightly deviated to the right along with an enlarged left cervical lymph node. Her complete blood count (CBC) was elevated at 19.7 x10³/ul (reference range 4-10 x10³/ul) with 12% lymphocytes, 2% monocytes, and 83% neutrophils. A rapid antigen test for S. pyogenes or Group A Streptococcus was negative. A CT exam of the neck was performed and a peritonsillar abscess of 1 x 1.3 x 1.6 cm was identified. The abscess was drained resulting in 1 ml of yellow purulent fluid which was sent to the microbiology lab for culture. The following was Gram stain was prepared from the abscess material.

Fusobacterium necrophorum Gram stain

Discussion

The Gram stain of this abscess showed 4+ PMNs and 4+ small, pleomorphic gram negative bacilli. Anaerobic culture grew Fusobacterium necrophorum, identified by MALDI-TOF MS with a confidence score of 2.2. F. necrophorum is a non-motile, non-pigment forming, pleomorphic gram negative bacilli. It is a strict anaerobe that tests catalase negative, indole positive, and lipase positive on egg yolk agar. Anaerobic antibiotic disk testing for this organism shows susceptibility to kanamycin and colistin with resistance to vancomycin.

The two most clinically relevant species of Fusobacterium are F. nucleatum and F. necrophorum. Because they are strict anaerobes which are often not recovered in culture, Fusobacterium spp. are an under-recognized cause of disease. F. necrophorum colonizes the oral cavity, and like other colonizing anaerobes, it tends to cause infections near the mucosal surface where it resides. F. necrophorum most commonly causes pharyngitis, recurrent tonsillitis, and other odontogenic infections. In adolescents, 10% of tonsillitis that is not caused by S. pyogenes can be attributed to F. necrophorum. These infections can progress to septic thrombophlebitis of the internal jugular vein (Lemierre’s syndrome), bacteremia, and rarely F. necrophorum can cause abscesses throughout the body. Because it is an anaerobic bacterium, susceptibility testing is rarely performed on isolates of F. necrophorum. They are highly susceptible to β-lactam–β-lactamase inhibitor combinations, carbapenems, and metronidazole.

Lemierre’s syndrome was of great concern in our patient since it is most commonly observed in adolescents and young adults that were previously healthy, like our patient. Fortunately, CT scan of the neck showed no indication of thrombophlebitis in our patient. After drainage of the abscess, she felt much better and was able to tolerate liquids. The patient was discharged from the ED with a course of amoxicillin/clavulanate (augmentin). Upon follow up in ENT clinic she gave a more through history of 4-5 episodes of sore throat over the past year.

References

Manual of Clinical Microbiology, 11^th edition
Principles and Practices of Infectious Disease, 7^th edition

-Erin McElvania, PhD, D(ABMM), is the Director of Clinical Microbiology NorthShore University Health System in Evanston, Illinois.

Microbiology Case Study: Generalized Weakness after a Trip to the United Kingdom

Case History
A 68 year old man presents with complaints of generalized weakness that started 3 weeks after his trip to the United Kingdom. The patient endorses night sweats, myalgia, fever, headaches, decreased appetite, mild nausea but no vomiting or diarrhea. He denied any history of recent rashes or arthralgia. The patient lives in a tick endemic area in the Northeastern part of the United States. Approximately 2 months ago he noticed an engorged tick while in the shower for which he completed a prophylactic course of doxycycline. He has sufficient titers for Lyme antibody; however no antibodies were present for Anaplasma. Blood smears were ordered for review.

Lab Identification
On the thin blood smears, there were multiple vacuolated, pleomorphic, ring-form like organisms seen in multiple infected red blood cells. No extracellular organisms were identified.

The organism was identified as a species of Babesia with 0.8% parasitemia and confirmed by PCR as Babesia microti.

Discussion
Babesia species are infectious protozoa which infect and cause lysis of red blood cells. Symptoms develop over the course of weeks to months and vary in severity. The most common symptoms are nonspecific flu-like symptoms (e.g., fever, chills, body aches, weakness, fatigue). If left untreated patients can develop hemolytic anemia, thrombocytopenia, disseminated intravascular coagulation, hemodynamic instability, and possibly death.¹

The main agents of human babesiosis are B. microti in the northeastern and upper midwestern regions of the United States, B. duncani in the western regions of the United States, and B. divergens in Europe. Occasionally on blood smears Babesia parasites can be difficult to differentiate from Plasmodium falciparum; Babesia species however are pleomorphic, vacuolated, and can appear inside red blood cells or outside red blood cells whereas Plasmodium falciparum are typically only seen inside red blood cells.¹ In most cases the diagnosis for babesiosis can be made on the basis of morphological features on thick and thin smears. However, for patients with subclinical symptoms, very low parasitemia with undetectable organisms on blood smears and a high clinical suspicion for babesiosis serologic and molecular testing can be offered. Serologic testing has 88-96% sensitivity and 100% specificity in patients with no concurrent history of malarial infections. There is some possibility of cross-reactions in serum specimens in patients with malarial infections.² Molecular techniques such as PCR are excellent tools for the purposes of screening and in addition can help to differentiate amongst the different variants of Babesia species.³ As in this case PCR was used to rule out possible infection with B. divergens due to the patient’s recent travel history to the United Kingdom. The two major antimicrobial regimens for babesiosis are atovaquone plus azithromycin for mild infection or quinine plus clindamycin for more severe infections.¹

References

Resources for Health Professionals. (2013, July 19). Retrieved August 21, 2017, from https://www.cdc.gov/parasites/babesiosis/health_professionals/index.html#dx
Krause PJ, Telford S RI, Ryan R, et al. Diagnosis of babesiosis: Evaluation of a serologic test for the detection of Babesia microti J Infect Dis 1994;169:923-926.
Hojgaard A, Lukacik G, Piesman J. Detection of Borrelia burgdorferi, Anaplasma phagocytophilumand Babesia microti, with two different multiplex PCR assays. Ticks and Tick-borne Diseases 2014 (5):349–351.

-Noman Javed, MD is a 1^st year anatomic and clinical pathology resident at the University of Vermont Medical Center.

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

Microbiology Case Study: A 53 Year Old Female with Extreme Dysphagia

Case History

A 53 year old Caucasian female presented to the emergency department with extreme dysphagia and pain in her mouth and throat. Her past medical history was significant for gastric adenocarcinoma for which she underwent a resection and received chemotherapy & radiation treatment. She had been hospitalized previously due to radiation esophagitis. On physical examination, she was cachectic but her vital signs were normal. Numerous ulcers were observed on her tongue and buccal mucosa. Her count blood count revealed she was pancytopenic with a white blood cell count of 0.19. An infectious disease work up was initiated and included blood and throat cultures as well as viral cultures of the oral ulcers for herpes simplex virus.

Laboratory Identification

capno1 — Image 1. Gram stain of a bacterial isolate that grew from blood culture showed long fusiform gram negative rods, which slightly tapered at both ends (100x oil immersion).

The blood culture was positive for long gram negative fusiform rods that tapered at both ends (Image 1). The organism grew as very small, whitish-yellow colonies on both blood and chocolate agars after 24 hours incubation in 5% CO₂ at 37°C. Biochemical reactions for both catalase and oxidase were negative. The isolate was identified by addition biochemical reactions by the reference bench as Capnocytophaga sputigena. Her throat and viral cultures were negative for additional pathogens.

Discussion

The Capnocytophaga genus is comprised of nine species that grow as facultative anaerobes and have a characteristic fusiform appearance on Gram stain. Organisms from this genus make up the normal flora of the oral cavity of humans as well as the oral microbiota of dogs and cats. Capnocytophaga spp. contribute to periodontal disease in adolescents and adults and the majority of disseminated infections arise from this endogenous source. The individuals at most at risk for septicemia include those that are immunocompromised (mainly neutropenic patients), alcoholics, intravenous drug users or those that lack a spleen.

In the laboratory, Capnocytophaga spp. is often first recognized by its characteristic Gram stain which shows long, fusiform gram negative rods that taper at both ends. Organisms with similar appearing Gram stain morphology include Fusobacterium spp. and Leptotrichia buccalis, but both of these bacteria exhibit anaerobic growth in contrast to Capnocytophaga spp which grows aerobically. Capnocytophaga isolates tend to grow slowly and require enriched media and increased CO₂ concentrations. The Capnocytophaga genus can further be broken down into a catalase- and oxidase-negative group and a catalase- and oxidase-positive group. Species in the first group include C. sputigena, C. gingivalis and C. granulosa. A notable species in the latter group includes C. canimorsus, which when it causes infection in humans it is most likely due to bites or contact with healthy dogs (25% colonization rate) or cats (15% colonization rate). Species differentiation can be challenging as some automated identification instruments can only identify to the genus level and many labs may not offer extensive biochemical work ups. However, the databases for the Bruker and Vitek MALDI-TOF MS currently include many of the species listed above.

In general, Capnocytophaga spp. are susceptible to broad spectrum cephalosporins, carbapenems, tetracyclines and fluoroquinolones. Resistance has been documented for aminoglycosides and colistin. In the case of our patient, her systemic infection was thought to be due to severe mucositis and the endogenous Capnocytophaga sputigena gained access to her blood stream via the numerous ulcers present. She responded well to antibiotic therapy and was discharged home.

Stempak

When Rapid Blood Culture Identification Results Don’t Correlate, Part 1: Clinical Correlation Needed

More and more laboratories perform rapid (i.e., multiplex PCR) blood culture identification. For the most part, it has been a wonderful addition to the laboratory workflow, not to mention the added benefits of provider satisfaction and improved patient care. Because the PCR only provides the organism identification (sometimes only to the family-level, i.e.; Enterobacteriaceae), laboratories must continue to culture the positive blood for definitive identification and/or antimicrobial susceptibility results. So what do you do when the results don’t correlate?

The Issue

From time to time, the PCR result is not going to correlate with the direct Gram stain or with the culture results. Although this is an issue one would fully anticipate, what do you do when this happens? Do you take some sort of action to arbitrate? Do you report the results as is?

First of all, the PCR assays do not detect all organisms. They only detect the most common bloodstream pathogens. Therefore, one should fully expect to observe cases in which the Gram stain would be positive, but the PCR results would be negative (scenario 1). This is not a surprise.

Additionally, one should also assume that the PCR will occasionally detect organisms that were present at the lower limit of detection of the Gram stain. An example of this would be that the Gram stain is positive for one morphology (i.e.; Gram-positive cocci), but the PCR is positive for two organisms (i.e.; Staphylococcus and a Proteus species). Most of these cases tend to correlate with culture. In other words, although the second organism was not originally observed in the Gram stain, it was detected via PCR and then it also subsequently grew in culture (scenario 2).

Another type of discordant result laboratories sometimes experience is when the organism detected via PCR does not grow in culture for whatever reason. Similar to scenario 2 stated above, except that the culture is also negative for the second organism (scenario 3). Perhaps the patient was treated with antibiotics and the organism is no longer viable for culture? Perhaps a sampling or processing error was to blame?

The Solution

Depending on the scenario and how much work you want to do, you can either repeat testing or try an alternative method. Take scenario 2 for example. If the PCR detects two organisms and the Gram stain is only positive for one, then review of the original Gram stain is warranted. It is possible that the Gram-negative was somehow missed. Our eyes tend to go to the darker, more obvious structures. Perhaps the Gram-negative organism was faintly stained and it was overlooked? It is also possible that the Gram-positive is present in much lower numbers and only Gram-negative organism was originally observed. If the Gram stain result remains the same after review (only one organism observed), then there is nothing much left to do except to wait for the culture. That being said, an alternative method, such as acridine orange can be utilized in this type of scenario (two different cell morphologies). Acridine orange is a fluorescent stain that improves organism detection, as it is more sensitive than the Gram stain (1, 2).

If only the Proteus is growing (and the Staphylococcus isn’t from scenario 2) and we normally subculture positive blood to blood, chocolate, and MacConkey agars, then perhaps including an additional media that inhibits Gram-negative growth would be beneficial.

Scenario 3 can be a little more difficult to solve because you can’t make a non-viable organism grow. It just is what it is. [Spoiler alert: in next month’s blog I plan to write about when you should change your thinking from true-positive to false-positive.]

Regardless of why the result is discrepant, our laboratory appends a comment to the discordant result which says, “Clinical correlation needed.” This lets the clinician know that the results are abnormal and that they must use other relevant information to make a definitive diagnosis. In addition to the comment, we also make sure the discrepancy is notified to laboratory technical leadership (i.e.; Doctoral Director, Technical Lead/Specialist). This allows us to keep track of discrepancies as they may become important to know about in the future (see next month’s blog).

The Conclusion

In terms of organism detection, nucleic assays (i.e., NAATs) can provide superior sensitivity over antigen and culture-based methods of organism detection (i.e., sensitivity = PCR > culture > Gram). From the laboratory perspective, other potential benefits of utilizing nucleic acid detection methodologies include decreased TAT, simplified workflows, and reduced hands-on time. In terms of patient care, many have noted improved outcomes due to increased sensitivity and decreased time to result.

Although advances in technology can significantly improve analytical performance, they can also add complexity to the post-analytical process. Making sense of the results can sometimes lead to confusion. It is important to know the product’s limitations and what your risk(s) is. This should already be known and included in your Individualized Quality Control Plan (IQCP). Lastly, guiding the clinician to proper result interpretation is also important to maintain valuable patient care.
References

Mirrett, S., Lauer, B.A., Miller, G.A., Reller, L.B. 1981. Comparison of Acridine Orange, Methylene Blue, and Gram Stains for Blood Cultures. J. Clin. Microbiol. 15(4): 562-566.
Lauer, B.A., Reller, L.B., and Mirrett, S. 1981. Comparison of Acridine Orange and Gram Stains for Detection of Microorganisms in Cerebrospinal Fluid and Other Clinical Specimens. J. Clin. Microbiol. 14(2): 201-205.

Martinez Headshot-small 2017

Microbiology Case: An Unusual Case of Cholangitis

Case History
A 64 years old male with a past medical history of atrial fibrillation, obstructive sleep apnea, and hypertension presented to the emergency room due to fevers and chills status post stent removal by endoscopic retrograde cholangiopancreatography (ERCP) 1 day earlier. The patient was admitted 6 weeks prior with Klebsiella bacteremia secondary to cholangitis due to an obstructive stone requiring ERCP with sphincterotomy and stent placement. In the emergency room the patient was febrile to 102.7F. Workup included an abdominal x-ray, right upper quadrant ultrasound, and CT abdomen and pelvis all of which were consistent with expected pneumobilia of the biliary tree due to his recent ERCP. On labs his lipase and liver function tests were within normal limits. Blood cultures were drawn and the patient was empirically started on piperacillin/tazobactam. Blood cultures flagged positive after 12 hours.

Laboratory Identification
Gram smear revealed gram negative bacilli. On the blood agar plates there were two different colony morphologies identified. Colony (A) was beta-hemolytic, oxidase positive, and white appearing on blood agar. Colony (B) was gamma-hemolytic, oxidase negative, and greyish appearing on blood agar. Both colonies were lactose fermenters on the MacConkey agar.

Kleb-Aero1 — Image 1. Gram stain from a positive blood bottle showing gram negative bacilli (100x oil immersion).

Image 2. Aerobic growth on blood agar showing two different colony morphologies. Colony (A) appears white with beta hemolysis and colony (B) appears grey with gamma hemolysis.

Image 3. Aerobic growth on chocolate agar showing two different colony morphologies. Colony (A) appears white and raised while colony (B) appears grey and flat.

Kleb-Aero4 — Image 4. Comparison of the two morphologically different colonies sub-cultured from blood agar to MacConkey agar. Both colonies (A) and (B) are lactose fermenters.

Using mass spectrometry, the MALDI-TOF positively identified the two organisms as Aeromonas species (colony A) and Klebsiella pneumoniae (colony B). The MALDI-TOF was unable to differentiate between A. hydrophilia and A. caviae species.

Discussion
Klebsiella pneumoniae is a known opportunistic pathogen implicated in nosocomial bacterial gastrointestinal infections. There are several proposed mechanisms by which this organism causes cholangitis which include ascension from the small bowel, contamination of the portal blood, or via translocation of the bowel wall following hematogenous seeding.¹

On the contrary, Aeromonas species are not native to the human gastrointestinal tract. These organisms are commonly found in freshwater and marine environments. They are gram negative, oxidase positive, facultative anaerobes. Most gastrointestinal infections caused by Aeromonas species are thought to be due to transient colonization of the GI tract and present asymptomatically or with mild diarrheal disease.⁶ Extra-intestinal wound infections are possible in the setting of a traumatic aquatic injury and cases of bacteremia have been reported; however these occur in the setting of malignancy or severe hepatobiliary disease.³

In the literature, there are 41 reported cases of hepatobiliary or pancreatic Aeromonas species infection. In almost all of these cases there are no documented aquatic environmental exposures. In one case series, 8/17 (47%) cases were due to nosocomial infections.³ One possible source for these infections can be the hospital water. Despite chlorination, Aeromonas species can be cultured from hospital water supply.⁴ Since many patients can be asymptomatic while transiently being colonized with Aeromonas species, it is possible that following an ERCP procedure, some organisms can be translocated from the GI tract to the biliary tree causing cholangitis.

To diagnose Aeromonas species a gram smear and biochemical testing should identify gram negative, rod shaped, non-spore forming, oxidase positive, glucose fermenting, facultative anaerobe organisms that are resistant to the vibriostatic agent O/129 and are unable to grow in 6.5% NaCl.² Their pattern of hemolysis on blood agar can be variable, although most species are beta-hemolytic. Mass spectrometry can further be used to identify at the level of the species. Most Aeromonas strains are resistant to penicillin and ampicillin and some automated MIC systems such as BioMeriuex Vitek may not be able detect the beta-lactam resistance.² Susceptibility studies should therefore be performed using standard agar dilution, broth microdilution, or using the Kirby-Bauerdisk diffusion method.⁷

Most Aeromonas species are susceptible to trimethoprim-sulfamethoxazole (TMP-SMX) and fluoroquinolones.⁵ There are some reported cases of fluoroquinolone resistance in patients that have a history of leech therapy. Aeromonas species can be isolated from the gut of the Hirudo medicinalis leech. These patients often receive systemic chemoprophylaxis to ciprofloxacin before undergoing leech therapy.⁵

References:

Kochar R, Banerjee S. Infections of the biliary tract. Gastrointest Endosc Clin N Am. 2013 Apr;23(2):199-218.
Morris, G.B., Horneman, A. (2017). Aeromonas Infections. UpToDate. Waltham, Mass.: UpToDate. Retrieved from uptodate.com.

Clark NM, Chenoweth CE. Aeromonas infection of the hepatobiliary system: report of 15 cases and review of the literature. Clin Infect Dis. 2003 Aug 15;37(4):506-13.

Picard B, Goullet P. Seasonal prevalence of nosocomial Aeromonas hydrophila infection related to Aeromonas in hospital water. J Hosp Infect 1987; 10:152–5.

Patel KM, Svestka M, Sinkin J, Ruff P 4th. Ciprofloxacin-resistant Aeromonas hydrophila infection following leech therapy: a case report and review of the literature. J Plast Reconstr Aesthet Surg. 2013 Jan;66(1):e20-2.

Gracey M, Burke V, Robinson J. Aeromonas-associated gastroenteritis. Lancet 1982; 2:1304–6.

Methods for Antimicrobial Dilution and Disk Susceptibility Testing of Infrequently Isolated Fastidious Bacteria. 3^rd ed. CLSI guideline M45. Wayne, PA: Clinical and Laboratory Standards Institute; 2016.

-Noman Javed, MD is a 1^st year anatomic and clinical pathology resident at the University of Vermont Medical Center.

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

Microbiology Case Study: A 73 Year Old Male with Fever, Lethargy, and Chills

Case History

A 73-year-old man presents to his primary care provider during the height of a bad influenza season with fever, lethargy, and chills. Symptoms started 24 hours prior to presentation. A rapid influenza rapid test was performed in the physician’s office and the result was negative for influenza A and B. What is the most likely cause of this man’s illness?

Answer

Influenza…but how can that be?

Discussion

Rapid antigen testing has been the mainstay for influenza testing since the 1980’s. These tests detect influenza A and B viral nucleoprotein antigens in respiratory specimens, giving a qualitative “positive” or “negative” result. Antigen testing was developed to shorten the turnaround time to results for common respiratory viruses influenza and respiratory syncytial virus (RSV), with an assay run time of approximately 15 minutes compared to the several days it takes for influenza detection by viral culture. Rapid antigen testing is very easy to perform, allowing CLIA-waived testing to be performed at point-of-care.

Unfortunately, rapid antigen testing has poor sensitivity. The most comprehensive analysis found the sensitivity of rapid antigen testing to be around 60% in adults and slightly higher (although still not good) in children. Due to the poor sensitivity, the CDC recommends only employing rapid antigen testing when the prevalence of influenza in the community is >10%…but why does the prevalence matter? Knowing the prevalence of a disease in your population allow you to calculate the positive and negative predictive value.

Positive and negative predictive values answer the question, “What is the chance that my positive test result means my patient has the disease (PPV) or what is the chance that my negative test result means my patient does not have the disease (NPV).” You can calculate the PPV or NPV of any assay by knowing the sensitivity and specificity of an assay along with the prevalence of disease in the community (Figure 1).

flu1 — Figure 1. Calculation of positive and negative predictive values.

Positive and negative predictive values fluctuate with the amount of disease seen in a community. For example, if testing for polio in the United States, where the virus has been eradicated, a positive test result by any method is far more likely to be a false-positive than a true-positive result. This is due to the low positive predictive value (PPV) of a positive test result in the setting of non-existent polio. The converse is true for negative predictive values (NPV). In the height of influenza season, a negative test result for influenza in a patient with signs and symptoms of influenza disease is more likely to be a false-negative than a true-negative result.

For influenza rapid antigen testing, the PPV is highest when influenza activity in the community is high (positive test result is likely to indicate influenza infection) and the PPV is lowest when influenza activity is low in the community is low such as in summer, when a positive influenza test result is most likely to be a false-positive result.

Conversely, NPV is highest when influenza activity is low in a community, and a negative test result is most likely indicating that the patient does not have influenza infection. NPV is lowest when influenza activity in a community is high, and a negative test result is more likely to indicate a false-negative result in a patient with influenza infection.

The specificity of rapid antigen assays is tied to the circulating influenza viral subtypes in a given season, and is generally quite high. Sensitivity and specificity do not change due to the prevalence of disease in the community, unlike positive and negative predictive values.

References:

Centers for Disease Control and Prevention (CDC) website on influenza testing (https://www.cdc.gov/flu/professionals/diagnosis/clinician_guidance_ridt.htm)
Altman Douglas G, Bland J Martin. Statistics Notes: Diagnostic tests 2: predictive values BMJ 1994; 309 :102
Chartrand C, Leeflang MM, Minion J, Brewer T, Pai M. Accuracy of Rapid Influenza Diagnostic Tests: A Meta-analysis. Ann Intern Med. 2012;156:500–511.doi: 10.7326/0003-4819-156-7-201204030-00403

-Erin McElvania, PhD, D(ABMM), is the Director of Clinical Microbiology NorthShore University Health System in Evanston, Illinois.