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

Case History

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

Image 1. Cyclospora cayetanensis.

Cyclospora cayetanensis

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

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

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

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

References

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

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

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

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

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

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

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

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

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

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

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

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

Microbiology Case Study: A 60 Year Old Male with Dysuria

Case Description

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

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

Discussion

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

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

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

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

How to Rock a Risk Assessment

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

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

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

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

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

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

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

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

Overview of Laboratory Tests for Cytomegalovirus

Introduction

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

Viral culture

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

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

Serology

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

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

Histopathology

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

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

pp65 antigenemia

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

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

Quantitative PCR

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

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

Conclusion

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

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

References

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

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

2019 Call for Abstracts

The Abstract Submission site is open for ASCP 2019. Last year, ASCP had a record number of submissions and we aim to bring in even more this year. ASCP’s selection process is known as highly competitive, and as a result, presenters receive close attention from big-name faculty and industry contacts. Some have even gained immediate funding opportunities to expand their research.

Abstracts can be submitted until 11:59pm CST on March 20th. Submit your abstract HERE.

Call for Abstracts 2019