Omicron: Variant of High Significance?

Omicron is now the dominant variant in the United States and gained that title faster than any variant before it. I have been tracking variants in the North Texas region since February of this year and detected the first Alpha variant (B.1.1.7). During this time, there were multiple substrains circulating. Some like Epsilon (origin California) rose in prominence then declined to extinction. Rise in Alpha (origin U.K.) and Delta variants (B.1.617.2, origin India) were tracked over the course of weeks, but Omicron has been tracked on a daily basis, since it is rising so quickly.

Many places are using S-Gene Target Failure (SGTF) as a surrogate for Omicron variant (Yale, University of Washington below).

Photo credit @NathanGrubaugh (Yale, Left) and @pavitrarc (UW virology, right)

SGTF occurs when the TaqPath COVID-19 multiplex test has 2/3 targets successfully amplify when the S-gene target does not or “drops out.”  This phenomenon was first observed in the Alpha variant, because the probe for this target overlapped a characteristic mutation: S:Del69_70 (deletion of the 69th and 70th amino acids in the spike protein from a 6 base pair deletion). This mutation is absent in Delta, but present in Omicron, so has been used as an early tracker of Omicron prevalence.

Most of this discussion is speculative and we won’t ever really know, but given the rate of transmission of this variant, it seems unlikely that it would have acquired so many mutations and not been detected before now. The most recent common ancestor is from over a year ago suggesting it was incubating for a long time.

We’ve seen a case of a person severely immunocompromised with no antibody response to vaccination + booster who still has an unmutated wild type strain in their system. With no immune pressure, the virus has not evolved.

However, in HIV+ patients with variable/ low immunity, there could be enough pressure to drive the immune evasion properties seen in Omicron. Southern Africa has over 30% of their HIV+ patients not on therapy who would be likely candidates for this type of host.

Did we see this coming?

Yes. Other immune evasive variants have arisen in areas with high prevalence of previous infection (Brazil/ S. Africa). Organisms evolve just enough to overcome the challenges in their environment. Thus the level of immunity provided by various immune exposures are approximately:

 Previous infection < 2x Vaccine < 2x Vaccine+ previous infection ~ x3 Vaccine

Scientists theorized that either Delta would evolve more immune evasive mutations or a totally new variant would arise. However, I didn’t think it would spread this quickly.

What is the impact?

Therapies. Most antibody therapies are directed as the business end of the spike protein—the receptor binding domain (RBD). The rest of the protein is covered in glycosylation modifications that block much recognition. Thus with many mutations in Omicron compared to the wild type strain (white), most therapeutic antibodies no longer bind/ inactivate viral replication.

Source: https://biorxiv.org/content/10.1101/2021.12.12.472269v1.full.pdf

Only one monoclonal antibody—Sotrovimab from GSK—is effective, because it binds a pan-coronovirus epitope outside of the RBD. However, this antibody is in short supply.

  • Thus, knowing which variant someone has can direct therapy. Several hospitals in our area are out of Sotrovimab, and only people with the Delta variant can access other options. Thus, knowing the variant in a short time frame has clinical implications.
  • Whole genome sequencing takes too long, so the FDA has agreed to review PCR genotyping approaches for clinical use. I have described some previous approaches, but many of these methods are useful as a screening method and would not have sufficient specificity to determine whether an omicron variant is present. Next time, I will discuss variant genotyping, why it is important, how it can be done, and what clinical actions can be taken with the knowledge.

Severity. There are signs that it is less severe. Is this due to increase in immune tolerance? We now have been prepared by either previous infection or vaccination to be protected from hospitalization or severe disease.

@Jburnmurdoch https://twitter.com/jburnmurdoch/status/1478339769646166019/photo/1

Or is the decline in severity due to lower pathogenicity? A recent non-peer reviewed study indicates the virus replicates x70 faster than Delta in the upper airways (left), but infiltrates cells 10% as well as the original strain.

From: https://www.med.hku.hk/en/news/press/20211215-omicron-sars-cov-2-infection?utm_medium=social&utm_source=twitter&utm_campaign=press_release

We all hope this will continue to be better news about the severity of Omicron, but from the lab side, I’ve heard of positivity rates >50% at some places. So this can still have a broad impact.

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

Microbiology Case Study: A 36 Year Old Male Traveler with Fever

Case Description

A 36 year old male presented to the emergency department with complaints of fevers, chills, night sweats, nausea, diarrhea, weakness, and decreased appetite for 6 days. He often travels between India and Dallas, and five months prior to presentation returned from two years abroad. While overseas, he developed similar symptoms, but due to COVID-19 restrictions, he was unable to see a provider at that time. His family doctor prescribed a course of medication for presumed malaria, which he completed but could not recall the name of the medication. He endorsed being ill for two weeks at that time and improved with medication to complete resolution of his symptoms. Prior to presentation, he also endorsed 3-4 episodes of non-bloody diarrhea per day and remembered a period of self-resolving chills a month prior. His fever and rigors were cyclic, occurring every other day, worsening up to presentation.

Given his travel history and symptomology, blood was drawn in the emergency department for analysis including a malaria smear. CBC and CMP were significant for elevated bilirubin (Total bilirubin 1.6 mg/dL, Direct bilirubin 0.4 mg/dL), leukopenia (3.60 x 10(9)/L), macrocytosis (92.5 femtoliters), thrombocytopenia (86 x 10(9)/L), and elevated CRP (6.6 mg/dL). His blood differential was significant for neutrophilia (91%), lymphocytopenia (7%), and monocytopenia (1%). The malaria smear was positive, and the patient was given a dose of artemether/lumefantrine in the emergency department. Plasmodium vivax was identified at a parasitemia of 0.5% (Figure 1), and the infectious disease service recommended admission for further workup including testing for G6PD deficiency prior to starting primaquine. He was not G6PD deficient, and an ultrasound of the spleen was unremarkable. The patient was treated with chloroquine for the erythrocytic and primaquine for the exo-erythrocytic stages of P. vivax malaria.

Figure 1. Photomicrograph of Plasmodium vivax ring forms observed in this patient’s Giemsa-stained peripheral blood smear, which are counted to determine the level of parasitemia in a patient’s bloodstream (500x oil immersion).

Discussion

Malaria is an infection caused by protozoan parasites of the genus Plasmodium. These organisms are transmitted by female Anopheles mosquitos upon taking a blood-meal. Human malaria is caused by five defined Plasmodium species: P. falciparum, P. vivax, P. ovale, P. malariae, and P. knowlesi.1 While not endemic to the United States, there is significant disease burden worldwide. In 2019, an estimated 230 million cases of malaria were reported causing approximately 409,000 deaths.2

The two lifecycles of Plasmodium sp. in the human host are classically defined as “erythrocytic” and “exo-erythrocytic”, involving red blood cells and hepatocytes, respectively. Plasmodium sporozoites are inoculated into the human host from the salivary glands of the mosquito upon feeding. From there, the sporozoites travel to the liver where they infect hepatocytes, mature into schizonts and ultimately merozoites. The infected hepatocyte then ruptures, releasing merozoites which enter the circulation and infect erythrocytes, initiating the erythrocytic cycle. This is a unifying trait of all Plasmodium sp. causing human malaria. Importantly, P. vivax and P. ovale form hypnozoites (dormant forms) in the liver, which can reactivate (oftentimes months to years later) following bloodstream clearance, resulting in relapse. It is therefore important that Plasmodium sp. infections be accurately speciated, as management of liver stage parasites differs from that of those in the bloodstream. By contrast, P. malariae and P. falciparum do not form hypnozoites and thus do not chronically infect the liver.

Plasmodium speciationis accomplished by evaluating thin and thick blood spears,4 allowing for assessment of parasite morphology and determination of parasitemia to guide patient management. In cases of P. vivax, the red blood cells are often enlarged (1.5 to 2 times the size of uninfected erythrocytes). Ring forms in all stages of development can be observed in P. vivax infection. These ring forms subsequently mature into trophozoites or gametocytes. P. vivax trophozoites exhibit a large, amoeboid cytoplasm, large chromatin dots, and fine yellow-brown pigment. Trophozoites subsequently develop into schizonts in the infected erythrocytes, subsequently rupturing leading to autoinfection. P. vivax schizonts are large with coalesced pigment and harbor 12 or more merozoites3 (Figure 2). P. vivax gametocytes are large and round to oval shaped and have scattered brown pigment, hemozoin, that may fill the erythrocyte (Figure 3). Gametocytes will migrate to the capillaries which are taken up by a mosquito upon taking a blood-meal, completing the Plasmodium lifecycle.

Figure 2. Photomicrograph of Plasmodium vivax merozoites in a schizont (1000x oil immersion) from this patient.
Figure 3. Photomicrograph of Plasmodium vivax gametocyte with malaria pigment (500x oil immersion) from this patient.

Here we present a case of relapsed P. vivax infection. Blood stage P. vivax parasites are susceptible to chloroquine, but dormant hypnozoites in the liver are resistant to its effects. Hypnozoites can be treated with primaquine, and thus routine management of either P. ovale or P. vivax usually consists of a combination of both antimalarial drugs. It is important to note that primaquine is contraindicated in cases of G6PD deficiency and pregnancy due to hemolytic complications,2 which is why this patient was tested prior to initiating therapy.

P. vivax has a worldwide distribution but has higher prevalence in colder climates as compared to other malaria species. P. vivax is most commonly encountered in Latin America and Southeast Asia. In addition to colder climate adaptation, P. vivax is interesting in that the parasite uses Duffy red cell antigens to enter erythrocytes and in populations with low frequency of Duffy on the surface of RBCs those groups are generally resistant to P. vivax infection. However, there have been rare cases of P. vivax in Africans who are Duffy-null.5

References

  1. Gladwin, M., Mahan, C. S., & Trattler, B. (2021). Malaria. In Clinical microbiology made ridiculously simple (pp. 343–346). essay, MedMaster, Inc.
  2. Menkin-Smith L, Winders WT. Plasmodium Vivax Malaria. [Updated 2021 Jul 23]. In: StatPearls [Internet]. Treasure Island (FL): StatPearls Publishing; 2021 Jan-. Available from: https://www.ncbi.nlm.nih.gov/books/NBK538333/
  3. Procop, G. W., Koneman, E. W., & Winn, W. C. (2017). Malaria. In Koneman’s color Atlas and textbook of diagnostic microbiology (pp. 1467–1470). essay, Lippincott Williams & Wilkins.
  4. Laboratory diagnosis of malaria: Plasmodium vivax. Laboratory Identification of Parasites of Public Health Concern. (n.d.). Retrieved September 14, 2021, from https://www.cdc.gov/dpdx/resources/pdf/benchAids/malaria/Pvivax_benchaidV2.pdf.
  5. Gunalan, K., Niangaly, A., Thera, M. A., Doumbo, O. K., & Miller, L. H. (2018). Plasmodium vivax infections of duffy-negative erythrocytes: Historically undetected or a recent adaptation? Trends in Parasitology, 34(5), 420–429. https://doi.org/10.1016/j.pt.2018.02.006

-Elisa Lin is a fourth-year medical student at UT Southwestern Medical Center in Dallas, Texas. She is interested in AP/CP track residencies.

-Clare McCormick-Baw, MD, PhD is an Assistant Professor of Clinical Microbiology at UT Southwestern in Dallas, Texas. She has a passion for teaching about laboratory medicine in general and the best uses of the microbiology lab in particular.

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

Microbiology Case Study: A Middle-Aged Man with Generalized Weakness

Case History

A middle age male with a past medical history of liver cirrhosis presented to the emergency department with one day of fever, chills, generalized weakness, and nausea. Complete blood count with differential showed leukopenia and neutropenia. Infectious work up was initiated including collection of 2 sets of blood cultures and imaging studies. A computed tomography (CT) scan of the abdomen revealed an irregularly shaped hypodense lesion in the right hepatic lobe concerning for abscess (Image 1). Ultrasound guided aspiration for the hepatic lesion yielded cloudy yellow bilious fluid, which was sent to the microbiology lab for aerobic and anaerobic cultures.

Image 1. CT scan of abdomen showing irregularly shaped hypodense lesion (yellow circle).

Two sets of blood cultures turned positive and Gram stain showed slender Gram positive rods in chains (Image 2). The aspirated fluid culture also showed 4+ Gram positive rods. Small gray colonies appeared on blood agar, chocolate agar, and Columbia Naladixic Acid (CNA) agar from both specimen types (Image 3). Lactobacillus rhamnosus was identified by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS). Minimal inhibitory concentration (MIC) was determined by broth microdilution assay and the organism was susceptible to penicillin while resistant to vancomycin. With appropriate antibiotics and abscess drainage, the patient’s condition improved and he was discharged to home.

Image 2. Gram stain of blood culture demonstrating gram positive rods in chains.
Image 3. Small gray colonies in blood agar from blood culture.

Discussion

Lactobacillus species are facultatively anaerobic, gram positive, non-spore forming rods that can have varying Gram stain morphology including short plump rods or long slender rods in chains or palisides.1 Lactobacillus species are bacterial inhabitant of the human mouth, gastrointestinal tract, and female genital tract. Isolation from clinical specimens could be considered by many to have questionable clinical significance.2 Lactobacillus species are often present in probiotics and fermented dairy products, including yogurt, and have been reported to provide benefits in gastrointestinal health,3–5 which led to increase in consumption of these products by many people, including our patient.

Cases of liver abscess and bacteremia caused by Lactobacillus species have been rarely reported in the literature and risk factors for the infection were immunosuppression, uncontrolled diabetes, hepatopancreaticobiliary disease, bacterial translocation, and use of probiotics or heavy dairy product consumption.6,7 The causative strains included L. rhamnosus, L. acidophilus, and L. paracasei.7

Pathophysiology of liver abscess and bacteremia due to Lactobacillus species is not well understood but it is postulated that several mechanisms may contribute to the pathogenicity of lactobacilli. Some strains are able to bind to intestinal mucosa, which may aid in translocation of the organism into the bloodstream. Also some strains can adhere to extracellular matrix proteins, aggregate platelets, and produce glycosidases and proteases.7 Furthermore, some strains are more resistant to intracellular killing by macrophages and nitric oxide.8

It is worth noting that many species of Lactobacillus are intrinsically resistant to vancomycin. However, they are usually susceptible to penicillin and ampicillin, as it was seen in our patient, and antimicrobial susceptibility testing can be performed by determining MIC of antimicrobials.9

References

  1. Goldstein EJC, Tyrrell KL, Citron DM. Lactobacillus Species: Taxonomic Complexity and Controversial Susceptibilities. Clin Infect Dis. 2015;60(suppl_2):S98-S107. doi:10.1093/CID/CIV072
  2. Chan JFW, Lau SKP, Woo PCY, et al. Lactobacillus rhamnosus hepatic abscess associated with Mirizzi syndrome: a case report and review of the literature. Diagn Microbiol Infect Dis. 2010;66(1):94-97. doi:10.1016/J.DIAGMICROBIO.2009.08.009
  3. Kligler B, Cohrssen A. Probiotics. Am Fam Physician. 2008;78(9):1073-1078. Accessed November 17, 2021. http://www.aafp.org/afp.
  4. Anukam KC, Osazuwa EO, Osadolor HB, Bruce AW, Reid G. Yogurt containing probiotic Lactobacillus rhamnosus GR-1 and L. reuteri RC-14 helps resolve moderate diarrhea and increases CD4 count in HIV/AIDS patients. J Clin Gastroenterol. 2008;42(3):239-243. doi:10.1097/MCG.0B013E31802C7465
  5. Adolfsson O, Meydani SN, Russell RM. Yogurt and gut function. Am J Clin Nutr. 2004;80(2):245-256. doi:10.1093/AJCN/80.2.245
  6. Omar AM, Ahmadi N, Ombada M, et al. Breaking Bad: a case of Lactobacillus bacteremia and liver abscess. J Community Hosp Intern Med Perspect. 2019;9(3):235. doi:10.1080/20009666.2019.1607704
  7. Sherid M, Samo S, Sulaiman S, Husein H, Sifuentes H, Sridhar S. Liver abscess and bacteremia caused by lactobacillus: Role of probiotics? Case report and review of the literature. BMC Gastroenterol. 2016;16(1):1-6. doi:10.1186/S12876-016-0552-Y/TABLES/1
  8. Asahara T, Takahashi M, Nomoto K, et al. Assessment of Safety of Lactobacillus Strains Based on Resistance to Host Innate Defense Mechanisms. Clin Diagn Lab Immunol. 2003;10(1):169. doi:10.1128/CDLI.10.1.169-173.2003
  9. CLSI. M45. Methods for Antimicrobial Dilution and Disk Susceptibility Testing of Infrequently Isolated or Fastidious Bacteria ; Proposed Guideline. Vol 35.; 2015. Accessed November 17, 2021. http://www.clsi.org.

Do Young Kim, MD is a medical microbiology fellow at University of Chicago (NorthShore). His academic interests include clinical microbiology and infectious diseases, epidemiology, and public health.

-Paige M.K. Larkin, PhD, D(ABMM), M(ASCP)CM is the Director of Molecular Microbiology and Associate Director of Clinical Microbiology at NorthShore University HealthSystem in Evanston, IL. Her interests include mycology, mycobacteriology, point-of-care testing, and molecular diagnostics, especially next generation sequencing.

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

Case History

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

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

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

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

Discussion

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

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

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

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

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

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

References

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

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

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

Peritoneal Problems

A 74 year old male patient with an extensive cardiac history initially presented to the ER with black stool, warranting a CT scan, upper endoscopy, and colonoscopy, identifying a large, obstructive mass in the colon, smaller, yet unresectable polyps, and subcentimeter liver lesions and lung nodules. The colonic mass was biopsied, consistent with adenocarcinoma; however, the liver lesions were too small to characterize. One month after the onset of symptoms, a right hemicolectomy was performed, and the pathology was signed out as moderately differentiated adenocarcinoma, microsatellite stable, with evidence of lymphovascular and perineural invasion, placing the patient’s stage at IIA (pT3, pN0, cM0). Through shared decision-making, the medical oncologist and patient elected for surveillance due to multiple comorbidities. Forgoing adjuvant therapy, the patient was discharged to physical therapy/rehabilitation. The patient returned for imaging 4 months after his hemicolectomy, demonstrating an enlargement in one of the liver lesions, but then, the patient was lost to follow-up for 20 months.


The patient reestablished care and surveillance imaging, which demonstrated a hypodense liver lesion (in a background of poorly visualized subcentimeter liver lesions), a nonocclusive thrombus in the right portal vein, a heterogenous enhancement of the left portal vein (suggestive of an underlying tumor thrombus), and an 8 cm heterogenous right adrenal mass. Based on the most recent CT scan, the differential diagnoses of the adrenal mass include metastatic disease or a primary adrenal lesion including adrenal cortical carcinoma or pheochromocytoma (for which biochemical analysis should be performed before attempting a biopsy). Extensive peritoneal lymphadenopathy was visualized as well. The area of the right hemicolectomy, however, did not show evidence of recurrence. After biochemical evaluation for metanephrines ruled out a pheochromocytoma, the patient underwent a CT scan-guided adrenal FNA and core biopsy.

The Diff-Quik smear assessed at the time of biopsy revealed a highly cellular specimen, some cells with bare nuclei, enlarged nuclei, and some pseudoglandular structures.

Images 1-2: Adrenal Gland, Right, Fine Needle Aspiration. 1-2: DQ-stained smears

Telepathology confirmed an adequate sample of tumor cells present, and core biopsies were obtained.

The following morning, the pap-stained smears and H&E cell block sections were screened. The cells appeared polygonal with a high N/C ratio and prominent macronucleoli. Cell arrangements formed thickened trabeculae. However, the cytoplasm is more granular than the lipid-rich cytoplasm seen in an adrenal cortical carcinoma. The H&E cell block sections depicted a beautiful trabecular pattern with endothelial cells wrapping the periphery.

Images 3-6: Adrenal Gland, Right, Fine Needle Aspiration. 3-4: Pap-stained smear; 5-6: H&E Cell Block sections.

The preliminary morphology was interpreted as carcinoma, and both cytotechnologist (or cytologist, as we now prefer to be called) and pathologist suggesting features of adrenal cortical carcinoma; however, the IHC markers proved otherwise!

Images 7-9: Adrenal Gland, Right, Fine Needle Aspiration, IHC Cell Block Sections. 7:HepPar1+; 8: Arginase+; 9: pCEA (canalicular pattern)+.

Other differential diagnoses considered renal cell carcinoma and pheochromocytoma (to be safe). The IHC profile ruled out adrenal cortical carcinoma as the cells of interest were negative for inhibin, calretinin, and Melan A. Negative PAX-8, EMA, AE1/AE3, and vimentin staining ruled out renal cell carcinoma, and negative chromogranin, synaptophysin, GATA-3, vimentin, and S100 staining enabled us to safely say that a pheochromocytoma was out of the equation as well. Positive staining for HepPar1, arginase, pCEA (canalicular pattern), and CAM5.2 supported the unlikely diagnosis of metastatic hepatocellular carcinoma (HCC).

This diagnosis placed the patient at Stage IV HCC. It came to light that the patient has a remote history of hepatitis and a high-risk history of drinking, contributing to a poor prognosis. Due to the patient’s condition, they held off on HCV antiviral therapy and decided to observing viral load through regular blood work. The patient and clinician discussed the risks and benefits along with alternatives of systemic therapy, as his multiple comorbidities still pose a significant risk. Immunotherapy was determined to be the best option to delay the progression of his cancer and maintain quality of life.

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

Validations/Verifications of Alternative Anticoagulants for Platelet Clumping

Platelet clumping can cause a falsely lowered platelet count on hematology instruments and can be difficult to resolve. With thrombocytopenia, physicians need an accurate count to diagnose, treat, or monitor patients. Clumping can be due to pre-analytic issues with specimen handling, can be caused by medications, or may be an in vitro phenomenon caused by anticoagulants. The clumping makes precise counting impossible and even estimates can be very tricky. If there are clumps, and recollection of the sample still yields platelet clumping, then many labs will have an alternate tube drawn or an alternative method to help resolve clumping.

Many of us have heard of using sodium citrate tubes for patients who have clumped platelets in EDTA. So, if you are having platelet clumping headaches, you can just order some sodium citrate tubes and start using those on your hematology analyzers, right? Not so fast. There are many published references of the use of sodium citrate tubes to resolve EDTA induced thrombocytopenia but we still see samples in which the clumping is not resolved with the sodium citrate tube. Published studies have shown that several other alternate methods have been helpful in resolving platelet clumping issues. These include drawing specimens in CTAD, ACD, or ‘ThromboExact’1 tubes, or adding amikacin or kanamycin to the EDTA after the specimen is drawn.

So, why can’t we just order one of these other tubes and start reporting results? Hematology analyzers are only FDA approved for EDTA tubes. Before you can use any modified method, and before you can report any patient results, your laboratory must do validation or verification studies to prove that the method produces valid results.

A validation provides objective evidence that a test performs as intended. A validation uses a defined process and is used when setting up and implementing a new test. One example is a laboratory developed test (LDT), which is a test performed by the clinical laboratory in which the test was developed. A LDT can be one that is neither FDA-cleared nor FDA-approved or can be one that is FDA cleared/approved but has been modified by the performing laboratory. The use of sample types or the use of collection devices not listed in manufacturer instructions constitute modifications, by this definition. In a validation, accuracy should be tested with at least 40 samples across the analytical measurement range (AMR). Correlations are then performed. Precision should be tested over approximately 20 days. A verification, on the other hand, uses an abbreviated process and is used when setting up and implementing new tests that are cleared or approved by FDA. Before reporting patient results, the laboratory must demonstrate that a test performs in agreement with prior claims and must demonstrate performance specifications are comparable to the manufacturer’s specifications. Verification therefore is a confirmation that a test method meets specified requirements and would be applied to a method which has already been validated. For a verification, a smaller sample size may be used, and precisions tested over 5 or more days.

Table 1. Validations vs. Verifications

So, which would you do if you wanted to use an alternate method for reporting platelet counts? Hematology analyzers are only FDA approved for platelet counts on EDTA, but the by which the sample is analyzed does not change with an alternate tube, so it may be possible to do a limited validation or verification with a smaller sample size. A laboratory needs to prove correlation, accuracy, and precision. Follow your laboratory SOPs for validation/verification and consult with your accrediting agencies, if necessary. A plan needs to be written and signed off by laboratory director. Choose the alternative method you wish to investigate and run correlations for platelet counts on EDTA and the alternate anticoagulant. If your instrument has more than one platelet mode, it is important to run samples in the mode which you would normally use for thrombocytopenia or flagged platelet counts. Apply any dilutional factors and calculate correlations. This data will be Included in your report, which, along with a procedure needs to be signed by the laboratory director.

The most important thing is to write a plan and a follow-up report according to your SOPs and to make sure any requirements of accrediting agencies are included. There can be some differences in interpretation of standards, so it is the laboratory’s responsibility to make sure what you have done meets the standards that apply to your lab.

The use of alternate tubes for platelet counts has been well reviewed in literature. Sodium citrate tubes are the most common, likely because they are the easiest to use and the most cost effective. Remember though that sodium citrate and other methods cannot resolve all case s of pseudothrombocytopenia. There are several special notes to consider. Counts from sodium citrate tubes are known to be stable for approximately 3 hours, after which counts decrease. As well, it has been shown in literature that sodium citrate tubes do show a negative bias. It has been reported that the 10% dilutional factor may be too low. Some studies have been done to determine dilution factors that correlate more closely with EDTA tubes, and researchers have suggested factor of 17%-25%. If your laboratory wishes to determine its own dilutional factor for sodium citrate or other tubes, this will also have to be included in your platelet studies. Lastly, CBCs are calibrated for EDTA, so only the platelet count should be reported from an alternative anticoagulant.

The end of another busy and challenging year is upon us, and at this time of year we can find ourselves rushed to finish ‘end of year’ tasks such as competencies and continuing education requirements. and a response to Sysmex’s recent webinar “Those Sticky, Tricky Platelets – Solving the Puzzle of Platelet Clumping” (Oct.20,2021). After the webinar I had many questions from techs asking, “Do we need to validate our alternative method?” and “How do we go about doing that?” The webinar discusses pseudothrombocytopenia and its causes in more detail than my earlier blog from Oct 2019, “Hematology Case Study: The Story of the Platelet Clump: EDTA-Induced Thrombocytopenia”. The webinar can be found at https://webinars.sysmex.com/webinars/11ae743e-ac99-47e7-acb7-2b24cedc1a1a and is available for CEU, free of charge.

References

  1. Baccini V, Geneviève F, Jacqmin H, et al. Platelet Counting: Ugly Traps and Good Advice. Proposals from the French-Speaking Cellular Hematology Group (GFHC). J Clin Med. 2020;9(3):808. Published 2020 Mar 16. doi:10.3390/jcm9030808
  2. Bizzaro N. (2013): Pseudothrombocytopenia. In: Platelets, Vol. 3, ed Bizzaro N, Elsevier, Amsterdam, pp. 989–997 
  3. Chae H, Kim M, Lim J, Oh EJ, Kim Y, Han K: Novel method to dissociate platelet clumps in EDTA-dependent pseudothrombocytopenia based on the pathophysiological mechanism. Clin Chem Lab Med 50, 1387–1391 (2012)
  4. Socha, Becky. Calibration and Calibration Verification: Who, What, Where, When, Why, How & Did I Pass or Fail?. AMT 81st Educational Program and annual meeting, 2019
  5. Zhou X, Wu X, Deng W, Li J, Luo W: Amikacin can be added to blood to reduce the fall in platelet count. Am J Clin Pathol 136, 646–652 (2011)
  6. https://www.cms.gov/Regulations-and-Guidance/Legislation/CLIA/downloads/6065bk.pdf
  7. https://www.cap.org/laboratory-improvement/proficiency-testing/calibration-verification-linearity
  8. https://www.westgard.com/cal-verification-criteria.htm
  9. https://labmedicineblog.com/2019/10/29/ hematology-case-study-the-story-of-the-platelet- clump-edta-induced-thrombocytopenia/
Socha-small

-Becky Socha, MS, MLS(ASCP)CMBBCM graduated 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 40 years and has taught as an adjunct faculty member at Merrimack College, UMass Lowell and Stevenson University for over 20 years.  She has worked in all areas of the clinical laboratory, but has a special interest in Hematology and Blood Banking. She currently works at Mercy Medical Center in Baltimore, Md. When she’s not busy being a mad scientist, she can be found outside riding her bicycle.

Microbiology Case Study: An Adult Patient with a Tender Mass and Rash

Case History

An adult patient with no significant past medical history presents with a tender right inguinal mass and rash over the right buttock measuring 5×7 cm. A skin punch biopsy was performed on the gluteal rash and sent to histopathology for analysis. Histology (Image 1) revealed an intradermal acantholytic vesicular dermatitis and associated folliculitis. Chronic inflammatory infiltrates surrounded neurovascular bundles as well as adnexal structures. Multinucleated Tzank cells were identified with the characteristic multinucleation, margination, and molding. Scattered eosinophilic Cowdry A inclusions were seen. Stains for bacteria and acid-fast bacilli (AFB) were not performed. A periodic acid-Schiff (PAS) stain (Image 2) demonstrated the absence of fungal elements.

Image 1. A hematoxylin and eosin (H&E) slide reveals a chronic inflammatory infiltrate surrounding (A) neurovascular bundles and (B) adnexal structures. (C) Tzank cells and (D) Cowdry A inclusions are also seen.
Image 2. A PAS stained slide of the same region as Image 1. (A) highlights a chronic inflammatory infiltrate where no fungal hyphae are seen.

Histopathology demonstrated “folliculitis suspicious for herpetic dermatitis.” PCR molecular testing for herpes simplex virus (HSV) and varicella zoster virus (VZV) were ordered on the punch biopsy. HSV was not detected; however, VZV was detected by PCR (Image 3, Image 4).

Image 3. The Simplexa VZV Direct Assay (Diasorin) targets a portion of the VZV DNA polymerase. The PCR amplification curve reveals the presence of VZV DNA (green) as well as that of the internal control (purple).
Image 4. A separate PCR assay targeting TP53 was performed to assess DNA quality of the fixed tissue. The presence of TP53 amplification in both the IC, the patient sample (Sample), as well as other samples on the same run (unlabeled) demonstrates the DNA quality is adequate. The absence of amplification of the NTC demonstrates a lack of nucleic acid contamination.

Discussion

Varicella zoster virus (VZV) is an enveloped double-stranded DNA virus belonging to the herpesviridae family.5 Transmission during primary infection occurs via inhalation of aerosolized respiratory secretions or lesional secretions, and to a lesser extent, via direct contact with lesional secretions. Transmission during secondary infection occurs mainly via physical contact with the secretions of herpetic lesions or the lesions themselves. The window for primary infection of transmissibility is 1-2 days before the onset of the rash lasting until either all lesions have crusted over or 24 hours have passed without the formation of new lesions, whereas secondary infections are only contagious during the presence of active lesions.6 Primary infection causes chicken pox, which is characterized by a vesicular rash, fever, and malaise. After primary infection, VZV resides in the dorsal root ganglia and trigeminal ganglia. VZV may reactivate, possibly as a result of stress or some other immunosuppressive state, as a painful vesicular rash known as shingles or herpes zoster. The rash is limited to the dermatome innervated by the ganglion from which the virus reactivated. Severe cases of shingles may result in meningitis, myelitis, as well as encephalitis, and can be fatal.1 Though the lesions of herpes zoster (secondary VZV infection) are infectious, they are significantly less so than those of varicella (primary VZV infection).6

The histology of VZV infection is characterized by intradermal and sub-epidermal vesicles with associated acantholysis, necrosis, and spongiosis. Tzanck cells demonstrate the characteristic “3 Ms” of multi-nucleation, marginated chromatin, and nuclear molding. The dermis is notable for perivascular, periadnexal, and perineural lymphocytic infiltrates. Folliculitis and syringitis may be present along with small vessel necrotizing vasculitis. Late stage lesions are notable for encrusted ulcers. Though there is significant histologic overlap between VZV infection as those caused by others in the herpes family, VZV histology tends to demonstrate a more substantial follicular involvement.2 Besides other herpes viruses, the differential diagnosis includes erythema multiforme, coxsackievirus, ecthyma contagiosum, pemphigus vulgaris and paravaccinia infection.3

While molecular methodologies are now the gold standard for diagnosis, a number of modalities including immunohistochemistry, immunofluorescence, in-situ hybridization, and serology can be used to aid in diagnosis.3 In the aforementioned case, diagnosis was made using a real time polymerase chain reaction (RT-PCR) assay (Simplexa VZV Direct Assay, Image 3) using previously extracted DNA. Forward and reverse primers target a well conserved portion of the VZV DNA polymerase. In between synthesis cycles, fluorescent probes anneal to the target sequence, separating the fluorophore from the quencher, thus generating a fluorescent signal. Amplification is measure by the cycle threshold (Ct), the number of PCR cycles needed for the fluorescent signal to exceed the background. An internal positive control (IC) is spiked in to assure negative results are not the result the presence of PCR inhibitors. To assess the quality of DNA present, a separate PCR was also performed on TP53, which amplifies if sufficiently high quality DNA is present, irrespective of the presence of VZV DNA (Image 4). A negative control (no template control, NTC) should be run to interrogate the presence of nucleic acid contamination.4

Treatment, if warranted, should be administered as soon as possible. Antiviral options include acyclovir, valacyclovir, or famcyclovir. Central nervous system, ocular, or renal VZV cases are considered emergencies and are typically treated with intravenous acyclovir.6 While resistance is rare, at least three mechanisms of resistance have been shown to endow VZV resistance to the aforementioned drugs: reduced or absent thymidine kinase, altered thymidine kinase activity leading to decreased phosphorylation of the drug, or decreased affinity of VZV DNA polymerase for acyclovir triphosphate.5, 8 If an infection with a resistant strain is identified or suspected, foscarnet is often used in place of acyclovir. Unlike the nucleoside analogs, this pyrophosphate analog does not rely on phosphorylation for the activation of its anti-VZV DNA polymerase activity.7 Historically plaque reduction assays were used, but this method is both labor intensive, low yield, and slow. Thus, molecular testing interrogating mutations in the DNA polymerase or thymidine kinase genes have increased in popularity.8

Two live attenuated vaccines are available, either in isolation or in combination with the measles mumps, and rubella vaccines (MMRV), in a 2 dose series to prevent primary infection. Since the VZV vaccine contains live virus, it should not be administered to pregnant women or the severely immunocompromised. Vaccine administration has been found to be 90% effective in preventing primary infection and 99% effective at preventing severe or complicated disease.7 Additionally, there is a recombinant vaccine consisting of the VZV glycophorin E protein in addition to an adjuvant that is used to prevent shingles. This formulation is recommended for adults over the age of 60 in prevention of secondary infections as well as to immunocompromised individuals at higher risk from exposure to the live attenuated vaccine.9

References

  1. Depledge DP, Sadaoka T, Ouwendijk WJD. Molecular Aspects of Varicella-Zoster Virus Latency. Viruses. 2018;10(7):349. Published 2018 Jun 28. doi:10.3390/v10070349
  2. Busam, K. J. Dermatopathology. 2nd Edition. Published 2014.
  3. Hall, B. Diagnostic pathology: Nonneoplastic Dermatopathology. 3rd Edition. Published 2021.
  4. Simplexa™ VZV Swab Direct REF MOL3655. 2021
  5. Sauerbrei A. Diagnosis, antiviral therapy, and prophylaxis of varicella-zoster virus infections. Eur J Clin Microbiol Infect Dis. 2016;35(5):723-734. doi:10.1007/s10096-016-2605-0
  6. https://www.cdc.gov/chickenpox/about/transmission.html
  7. https://www.cdc.gov/vaccines/vpd/varicella/hcp/index.html
  8. Piret J, Boivin G. Antiviral resistance in herpes simplex virus and varicella-zoster virus infections: diagnosis and management. Curr Opin Infect Dis. 2016;29(6):654-662. doi:10.1097/QCO.0000000000000288
  9. https://www.cdc.gov/vaccines/hcp/vis/vis-statements/shingles-recombinant.html

-Jeremy Adler, MD is a Molecular Genetic Pathology fellow at the University of Chicago Medicine and NorthShore University HealthSystem. He completed his MD at SUNY Stony Brook and his AP/CP residency at the Pennsylvania Hospital of the University of Pennsylvania Health System.

-Paige M.K. Larkin, PhD, D(ABMM), M(ASCP)CM is the Director of Molecular Microbiology and Associate Director of Clinical Microbiology at NorthShore University HealthSystem in Evanston, IL. Her interests include mycology, mycobacteriology, point-of-care testing, and molecular diagnostics, especially next generation sequencing.

Safety in the Moment

Often I am asked how one who is responsible for laboratory safety (yet has other duties as well) can get the job done well. In today’s labs there is tight staffing, tight budgeting, and a score of regulatory duties that must be accomplished, and not all of these things revolve around safety. Many who oversee the lab safety program also must run the point of care program, the lab quality program, or even manage all of the day to day operations of the department. It’s a great deal to juggle, but there are methods you can use to make sure that laboratory safety doesn’t take a back seat.

One way to incorporate safety into your multiple roles each day is to start every meeting or huddle with a safety moment or story. Ask for a team member to discuss a safety story they witnessed or in which they were involved. Placing safety first lets the team members know it has priority, and relating an issue or incident has benefits as well. The safety moment may be as brief as reporting on how an employee provided PPE to a vendor that came into the department. That is a safety success worth mentioning, and there are doubtless others that can be mentioned. These safety stories may also be those that do not necessarily illustrate a success. Telling people about an incident and asking how it could have been avoided is a fast yet educational plus for your safety culture. Reviewing safety incidents is also beneficial so that others know what happened and they can be thinking of how to avoid the same thing from happening to others or themselves. Talking about safety in these ways takes little time, but if safety is incorporated into the language of the department, the culture will remain improved, and it is easy to fit this habit into your schedule.

Acting as a consistent role model is another way to incorporate safety into your multiple roles. Make sure you wear the correct clothing and shoes. If you walk in and out of the department, you should dress the part. Open-toed shoes or mesh sneakers should not be worn. Wear PPE when performing any work in the lab, including huddles or team meetings. It doesn’t take any extra time to model the safety behaviors you expect from the staff, and doing this shows the staff where safety stands in the department.

A third way to insert safety into your busy day is to make sure you are able to quickly spot safety issues and address them immediately. Developing your “Safety Eyes” is a vital tool – learn how to notice safety problems as you work in the lab. Train yourself to be able to do this by looking for one thing each week. For instance, look for PPE and dress code issues on week one. Purposely notice what people are wearing on their feet, look for proper PPR like lab coats and gloves. Check to see that they are worn properly. If you do this for one week, you will become much better at noticing issues with just a glance. The next week look for proper chemical labels, then fire safety issues, etc. Once your Safety Eyes are enabled, you will be able to easily see issues and manage to rectify them while performing your other lab duties.

No matter your role in the laboratory, part of the job involves talking to other people. Make safety a part of those conversations when the opportunity arises. You might speak to your lead technologist about an instrument installation. Ask about new reagents that might need to be added to the chemical inventory.  Find out if there will be new waste streams generated. Was a risk assessment performed to look for other possible dangers?

Incorporating safety into your already busy day might seem like an impossibility, but it can be done. It is important that it is done. You are managing different parts of the lab, but if people are getting injured and exposed because there is no focus on safety, there won’t be much left to manage! Try these few ways to blend safety into your schedule- add one at a time and see how it works. In time you will notice that these small tasks make a big improvement on your lab safety culture.

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

Microbiology Case Study: A Female with Diabetes and Renal Disease

Case history

A middle-aged female with a past medical history of diabetes and end stage renal disease resulting in kidney transplant presented for evaluation of right hip and knee pain for the previous two months. An MRI of the hip revealed a large effusion with evidence of septic arthritis, myositis in the surrounding muscle, and osteomyelitis of the hip. Blood cultures remained negative for the duration of her presentation. The patient underwent a joint aspiration, and synovial fluid was sent to the microbiology laboratory for culture. Due to subsequent culture positivity and the extent of the involvement of the surrounding anatomy, the patient was started on ceftriaxone and underwent a total joint replacement. Her symptoms improved post-procedure, and post-operative vertebral MRI and TTE revealed no evidence of osteomyelitis or endocarditis. The patient was discharged on post-operative day six with continued IV ceftriaxone for an additional 5 weeks.

Laboratory identification

The synovial fluid received in the microbiology laboratory was plated onto blood, chocolate, and MacConkey agars. No organisms were visible on direct Gram stain, but the culture revealed scant growth of alpha-hemolytic colonies on blood and chocolate plates. These colonies were comprised of faintly staining gram positive rods (Image 1). The organism was catalase negative. Given the characteristic appearance by Gram stain, the organism was inoculated to a triple sugar iron (TSI) slant where it demonstrated H2S production. A definitive identification of Erysipelothrix rhusiopathiae was achieved by MALDI-TOF MS.

Image 1. Synovial fluid culture sent to the microbiology laboratory. E. rhusiopathiae colonies growing on Sheep’s blood agar are denoted by black arrowheads. Characteristic Gram stain of the E. rhusiopathiae colonies from the plate revealing poorly staining gram positive rods. TSI slant from the colonies demonstrating H2S production.

Discussion

Erysipelothrix rhusiopathiae is a facultatively aerobic, non-spore forming, gram positive pathogen that is a resident of the digestive and respiratory tracts of mammals, bird, fish, and pigs.1 It is the etiological agent of Swine Erysipelas, causing either an acute septicemia, cutaneous disease, endocarditis, or chronic arthritis in pigs. Human infections with E. rhusiopathiae are usually due to exposure to infected animals or contaminated animal products or environments. Certain occupations with frequent animal exposure are at increased risk for infection (including fishermen, veterinarians, farmers, and butchers). Infection requires entry into the skin through cutaneous abrasions, which can be caused by sharp hooks, fish scales, teeth, and other occupational tools or hazards that damage epithelial barriers.1,2

Human E. rhusiopathiae infection can manifest as three distinct forms. An acute, localized cellulitis named eryspieloid (not to be confused with streptococcal erysipelas) is the most common manifestation. This usually impacts the hands, fingers, or other parts of the upper extremities that have contact with animals or animal products.3 A generalized cutaneous form more often associated with systemic symptoms including fever, joint aches, lymphadenitis, lymphadenopathy, and arthritis can also occur. Finally, septicemia frequently associated with endocarditis is a third manifestation. E. rhusiopathiae endocarditis is often subacute, with a tropism for native valves (particularly the aortic valve). Due to its indolent nature, this presentation often requires valve replacement at the time of diagnosis and is associated with increased mortality.1,4 While cases of non-severe eryspieloid may self-resolve, ampicillin or penicillin are the treatments of choice for cutaneous and systemic infections. Cephalosporins and fluoroquinolones are also efficient alternative agents.3 Importantly, the organism is intrinsically resistant to vancomycin, thus accurate and timely identification is critical to ensure appropriate intervention (Image 2). Susceptibility testing is generally not performed but may be useful in the setting of penicillin allergy.

Image 2. E. rhusiopathiae is intrinsically resistant to vancomycin. E. rhusiopathiae exhibits elevated MICs to vancomycin. Penicillin is the treatment of choice.

Laboratory identification of E. rhusiopathiae can be challenging.  Erysiepelothrix can easily decolorize during gram staining and can be mistaken as gram negative due to lack of stain retention. Additionally, the cells can exhibit variable morphologies including pairs, chains, and filaments. Colonies can also exhibit variable morphotypes when grown on routine media, including both rough and smooth forms.2An environmental exposure to animals was investigated in this patient’s case to possibly serve as the source of infection. While a direct link cannot be definitively proven, it was revealed that the patient owned a large fish tank which she regularly cleaned which could have been a potential source of infection. 

References

  1. Wang Q, Chang BJ, Riley TV. 2010. Erysipelothrix rhusiopathiae. Veterinary Microbiology 140:405-417.
  2. Clark AE. 2015. The Occupational Opportunist: an Update on Erysipelothrix rhusiopathiae Infection, Disease Pathogenesis, and Microbiology. Clinical Microbiology Newsletter 37:143-151.
  3. Veraldi S, Girgenti V, Dassoni F, Gianotti R. 2009. Erysipeloid: a review. Clinical and Experimental Dermatology 34:859-862.
  4. Brooke CJ, Riley TV. 1999. Erysipelothrix rhusiopathiae: bacteriology, epidemiology and clinical manifestations of an occupational pathogen. Journal of Medical Microbiology 48:789-799.

-Timothy J. Kirtek, M.D., originally from Grand Blanc, Michigan, graduated from American University of the Caribbean School of Medicine located on the island of Sint Maarten. There, he conducted research on tropical arboviruses including Dengue, Chikungunya, and Zika viruses. He then returned to Michigan to complete his clinical training and, upon graduation from medical school, moved to Dallas, Texas where he is currently an Anatomic and Clinical Pathology resident physician at UT Southwestern.

-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.

Where have all the Techs Gone?

Electronic media is replete with articles and editorials of employers lamenting the shortage of workers. Signs offering hiring bonuses hang outside of restaurants, stores, and other retail outlets all across the country.

The inability to find workers has forced employers to take another look at their business model and reevaluate whether the model is still viable in its current form. The power balance in the employer/ employee dynamic has shifted. Employers accustomed to having their choice of applicants now find themselves scrambling to find workers.

No schools, No students

The healthcare industry, including the medical laboratory, is not exempt from the shortage despite healthcare experts and administrators knowing that the trending laboratory employee shortage was inevitable years ago.

Laboratory school administrators and managers have been sounding the alarm about the lack of community college and university medical technology program applications. Many academic medical technology programs are shuttered due to a lack of students.  The decrease in the number of students going into the laboratory field and the normal attrition rate of older workers retiring or moving on to higher-paying occupations has led to a high vacancy rate and a loss of expertise.

Burnout

The pandemic has added more pressure on a cohort of employees experiencing the stress of a new and unknown danger. These allied health professionals were (and are) the front-line response to a disease threatening everyone, regardless of economic or social demographics. Lab worker burnout has become a documented phenomenon

We call them heroes, but in reality, these are the same people working every day (pandemic or not), serving patients and delivering quality test results. Labs across the nation are filled with these everyday people. But just like everyone, laboratory workers have families, feelings, and needs they are trying to meet while being asked to give a little more. Many have little left to give and are now leaving the field to pursue other less stressful occupations or to simply enjoy the life they have worked so hard to build.

Start recruiting early

How can healthcare organizations stem the tide of those choosing to leave the lab and simultaneously attract young fresh minds to the unglamorous and less financially rewarding but necessary field of laboratory testing?

Presentations to elementary school children are a great way to introduce the next generation to the laboratory field. What child doesn’t like looking into a microscope to see their own red and white blood cells? Roadshows put on in junior high and high schools are a great way to kindle interest in healthcare just when students are beginning to ponder the question of what they want as a career.

Educational Aid

The cost of college continues to rise. Scholarships are often garnered by high-performing “A” students. But there is a pool of “B” students that could also benefit from financial assistance and would be just as welcomed into clinical laboratories. Broadening and diversifying the qualifications to receive a scholarship and financial aid could conceivably add to the pool of potential laboratory workers. Another unique idea is to allow laboratory workers’ dependents access to unused employee educational benefits.

Wellness in the Lab

Resources should also be dedicated to retaining technicians and technologists who are considering leaving the laboratory field.  The level of compensation is meaningful, but studies have shown that employees often leave the job for more esoteric reasons. Reducing stress, supporting a culture of wellness, inclusiveness, and belonging can differentiate one workplace from another. The theme of workplace wellness was extensively discussed at this year’s ASCP 2021 annual meeting in Boston.

The Need is Real

The pandemic has highlighted the importance of the laboratory to the health of the nation. The medical laboratory should use this moment in the spotlight to advocate for more resources and emphasize the necessity for more laboratory programs and students to meet the future testing needs of the nation.

Of course, many lab managers are wondering what to do today to stem the slow leak of personnel. Providing mental health support and financial incentives do work to keep these knowledgeable workers in the lab. Managers realize that laboratory science is a demanding high acuity job with little or no margin for error. To maintain quality, the healthcare industry will need to change its perceptions about the laboratory and address the lack of technicians and technologists with the same interest and retention resources given to nurses and doctors.

-Darryl Elzie, PsyD, MHA, MT(ASCP), CQA(ASQ), has been an ASCP Medical Technologist for over 30 years and has been performing CAP inspections for 15+ years. Dr. Elzie provides laboratory quality oversight for four hospitals, one ambulatory care center, and supports laboratory quality initiatives throughout the Sentara Healthcare system.