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

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

The Issue

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

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

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

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

The Solution

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

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

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

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

The Conclusion

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

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

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

 

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

Time Mastery

One of my favorite song lyrics is from “When I Find Home” by Cody Chestnutt: “I only got time to think about the time I don’t have” I like this lyric, because when I get really busy I sometimes enter this “freeze” moment, where I am stuck thinking about all the things I have to do without being able to do any of them.

As our work continues to get busier and busier, it is becoming more critical to have good time management skills. However, to actually master time, people need more than To-Do lists. This course focuses on twelve different categories of time mastery and participants assess their skill level in each of these areas:

  1. Attitudes
  2. Goals
  3. Priorities
  4. Analyzing
  5. Planning
  6. Scheduling
  7. Interruptions
  8. Meetings
  9. Written Communication
  10. Delegation
  11. Procrastination
  12. Team Time

However, not all of the categories are equally important in a current position. I might have no direct reports, so even though I might score low in that category, it is not really important in my current job. Based on participants’ answers, the assessment automatically creates a Skills Gap Analysis, a table in which the categories are organized according to two axes: less important to important and less skill to more skills. These tables gives participants a quick overview of which categories they have marked as more important, but have less skill in. In other words, these are the areas of development.

Mastering time and moving away from thinking about the time that I do not have, has allowed me be more proactive about my time and schedule. My written communication and meetings are more productive and better organized; I am clearer in my delegation and define authority levels; I follow my yearly goals more closely and I take the time to analyze when and why I am interrupted or interrupting, to understand what could have been communicated better. Mastering time has allowed me to rarely experience stress while at the same time being more productive. I only got time to think about the time I do have.

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-Lotte Mulder earned her Master’s of Education from the Harvard Graduate School of Education in 2013, where she focused on Leadership and Group Development. She’s currently working toward a PhD in Organizational Leadership. At ASCP, Lotte designs and facilitates the ASCP Leadership Institute, an online leadership certificate program. She has also built ASCP’s first patient ambassador program, called Patient Champions, which leverages patient stories as they relate to the value of the lab.


 

When many of us are asked if we would be interested in learning more about improving our time management skills, our response, maybe “Yawn, I got this, after all, I am a busy professional.” What we often don’t realize is that getting to the goal, no matter how hurried or rushed, is not just often about you. Our work pace impacts others in the workplace and if we want to become a true leader, we need to master ourselves first. The first place to start on this is how we manage our time.

We have an expression in our family: “we’re always working for the farm.” When you are running a farm, you don’t get to choose your own timing or schedule in the projects that must be done. The seasons come and go and there is planting and harvesting to be done on nature’s time. There are animals to be fed and fences to be mended on nature’s time. Understanding some of these basics help us to realize that there is time in one’s schedule that we can control and time in one’s schedule that we don’t control.

As for the time you can control at work, you can select when to answer your emails, and when to have “that open door” for workplace issues, and when and how to prioritize your projects. In some cases, like the farm, you will not get to always control your own schedule and choose when important events or meetings happen. So to help with this, we can paraphrase Mark Twain: “Eat your frogs early.” This means doing your biggest and hardest task first thing every day so that you prevent procrastination and free up time in case other urgent situations emerge. For some, that may be a phone call dealing with a patient or employee complaint; for others, it may be tackling an unresolved operational issue that needs to be urgently addressed. Whatever it is, go at it first and efficiently and get the job done.

When we look to improve how we master time, we need to have an understanding of what is urgent and important and what is important, but not urgent. The best advice for time management is to work more on what is important, but not urgent, to prevent everything from becoming a last minute urgent need. If you are often focused on urgent issues every day, you are simply putting out the fires at work and never getting to the optimal operational efficiency in your area. You can begin to master this by simply blocking out a time every day that you will work on these important projects. This will soon become part of your habitual schedule and that job will get done over time by breaking these projects up into smaller blocks of time. The most productive writers often say that they sit down with their computer to write during a certain time of day, whether they feel like it or not, and much to their surprise they are able to make progress. Yes, this even works for the great story tellers of our time, such as Earnest Hemingway, who sat down every morning at the same time to write.

One of the best ways to become a time master is to understand your own biases, strengths and weakness about time management. Are you good at delegating tasks that can be performed by others? Do you lose track of time when you are interrupted in your office? There are tools that you can use to assess your time management skills, and help you work to develop better habits for improved productivity and better balance. As you begin to become more proficient in time management, you will find that your overall work place and life stress will also decrease, as you find more “time” to take on more of those projects that bring balance and joy into your work and life.

 

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-Dr. Deborah Sesok-Pizzini, MD, MBA is a Clinical Pathologist at the University of Pennsylvania, Perelman School of Medicine, who specializes in Blood Banking and Transfusion Medicine at the Children’s Hospital of Philadelphia.  She has a strong interest in resident leadership development, patient safety and quality, and is currently serving as a member on the ASCP Fellow Council.  She is a graduate of the ASCP leadership institute certification program and has an MBA from Villanova University with a concentration in finance.  

Microbiology Case: An Unusual Case of Cholangitis

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

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

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Image 1. Gram stain from a positive blood bottle showing gram negative bacilli (100x oil immersion).
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Image 2. Aerobic growth on blood agar showing two different colony morphologies. Colony (A) appears white with beta hemolysis and colony (B) appears grey with gamma hemolysis.
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Image 3. Aerobic growth on chocolate agar showing two different colony morphologies. Colony (A) appears white and raised while colony (B) appears grey and flat.
Kleb-Aero4
Image 4. Comparison of the two morphologically different colonies sub-cultured from blood agar to MacConkey agar. Both colonies (A) and (B) are lactose fermenters.

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

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

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

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

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

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

References:

  1. Kochar R, Banerjee S. Infections of the biliary tract. Gastrointest Endosc Clin N Am. 2013 Apr;23(2):199-218.
  2. Morris, G.B., Horneman, A. (2017). Aeromonas Infections. UpToDate. Waltham, Mass.: UpToDate. Retrieved from uptodate.com.
  1. Clark NM, Chenoweth CE. Aeromonas infection of the hepatobiliary system: report of 15 cases and review of the literature. Clin Infect Dis. 2003 Aug 15;37(4):506-13.
  1. Picard B, Goullet P. Seasonal prevalence of nosocomial Aeromonas hydrophila infection related to Aeromonas in hospital water. J Hosp Infect 1987; 10:152–5.
  1. Patel KM, Svestka M, Sinkin J, Ruff P 4th. Ciprofloxacin-resistant Aeromonas hydrophila infection following leech therapy: a case report and review of the literature. J Plast Reconstr Aesthet Surg. 2013 Jan;66(1):e20-2.
  1. Gracey M, Burke V, Robinson J. Aeromonas-associated gastroenteritis. Lancet 1982; 2:1304–6.
  1. Methods for Antimicrobial Dilution and Disk Susceptibility Testing of Infrequently Isolated Fastidious Bacteria. 3rd ed. CLSI guideline M45. Wayne, PA: Clinical and Laboratory Standards Institute; 2016.

 

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

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

Packing for the Trip

If you are sending specimens outside of your lab for testing or for other purposes, there are many things to consider. Not just anyone in the laboratory should prepare the specimens for shipment, specific training is required. Staff needs to have knowledge about packing procedures, specimen safety considerations, and how to fill out the necessary paperwork.

The safe transport of laboratory specimens is regulated by at least four different agencies. The United States Department of Transportation (DOT) generates specimen packaging and shipping regulations and updates them every October. They become effective on April first of the following year. The International Civil Aviation Organization (ICAO) provides technical instructions for domestic and international air shipments. Dangerous Goods regulations are published by the International Air Transportation Association (or IATA). These annual updates take effect every first of January, and they comply with ICAO instructions as well. The fourth regulatory body affecting specimen shipping is the United States Postal Service (USPS). The USPS regulations are synchronized with those of both the DOT and IATA. In fact, most of these regulatory agency’s shipping regulations are similar because they have been harmonized with the United Nations Model Regulations.

Dangerous Goods are categorized under classes and divisions, and laboratory specimens fall under Class 6, Division 2; Infectious substances. For the purposes of shipping, these infectious substances are further divided into Category A (specimens known or reasonably expected to contain pathogens) and category B (human or animal specimens which do not contain pathogens). Very specific training for lab staff is required for those who will package and ship these types of products. A laboratory that sends such specimens is considered the shipper, and shippers are viewed in the eyes of the law as responsible for the package until it reaches its final destination. That responsibility involves proper and regular training, specific packaging instructions, and management of shipping paperwork.

Dangerous Goods packing and shipping training includes General Awareness information. This includes an overview of the various regulations surrounding specimen shipment as well as enabling the trainee to recognize and identify the hazardous materials that fall under the regulations. Function-Specific training is general instruction on how to package dangerous goods, how to properly label parcels, and how to fill out the required paperwork. Training shippers about Safety is also required. Staff must be given information about the hazards associated with handling dangerous goods, and there must also be training given about how to handle emergencies such as accidents or spills. Lastly, Security training is required because the materials being shipped could be misused in a way that could cause harm to others (i.e. a terrorism event).

The paperwork that must be filled out and that must accompany the package is called a waybill or a shipper’s declaration. A lab must have two original completed and signed forms. Two of the signed forms travel with the shipment to its final destination, and one will be retained by the carrier. There are very specific instructions about how to fill out a shipper’s declaration, and only those trained should do so.

Not all specimens and chemicals shipped by a lab are considered Dangerous Goods, and they may not fall under the regulations for labs that transport them. The DOT designates some items as “Materials of Trade,” and they are not regulated under transport laws. Material of Trade are those items which are carried on a motor vehicle to directly support a principal business of a private motor carrier (such as a private courier). That means that many diagnostic specimens and some hazardous chemicals (under 8 gallons depending on the chemical) may be transported without shipping papers, emergency response plans, and specialized training.

Proper training for those shipping Dangerous Goods takes time, and the information included in the training is much more in-depth than what has been discussed here. Those who are trained should be tested on the information taught, or they can provide a demonstration of proper packaging and paperwork management. Each successful trainee should be given a certificate of completion, and that record needs to be retained for at least 36 months. Training should be repeated every two years in order to satisfy the requirements of all regulatory agencies.

Laboratories across the country package and send diagnostic specimens for testing or for other purposes. Sometimes, depending on how and where those specimens will be transported, very specific regulations will apply, and specialized staff training will be required. Before you get those samples packed and ready for their trip, make sure your lab is following the regulations that will keep your staff (and those who will transport and receive the packages) safe from harm.

 

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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.

Hematopathology Case Study: A 64 Year Old Man with Widespread Lymphadenopathy

Case history

A 64-year-old, previously healthy man presented with a history of cervical and axillary lymphadenopathy of unknown duration. He did not endorse night sweats, weight loss, or fever. Radiologic examination (CT chest and MRI abdomen) revealed numerous enlarged mediastinal, peritracheal, periaortic, periportal and retroperitoneal lymph nodes. He underwent excisional biopsy of a 3.5 cm axillary lymph node.

foll-lymph

Microscopic Description

Histologic examination of the node revealed distortion of nodal architecture by a proliferation of neoplastic-appearing follicles. Follicles were distinct from one another, and closely packed. In areas the follicles were present back-to-back. Follicular centers were comprised of mostly small, cleaved centrocytes and showed no obvious zonation. There was loss of tingible body macrophages.

Immunophenotyping

Immunohistochemical analysis revealed CD20-positive B cells in a follicular pattern. The germinal centers revealed an underlying follicular dendritic meshwork highlighted by staining for CD21. Interestingly, while the germinal centers demonstrated immunopositivity for BCL-6, there was minimal to absent CD10 staining on follicular B cells. Analysis of BCL-2 staining revealed only few cells to be positive within the follicular centers, consistent with resident follicular helper T cells (Th cells). Equivalent numbers of CD3 and CD5 positive T cells were noted in the interfollicular zones. The Ki-67 proliferation index was estimated at 15-20% within follicular centers. Flow cytometric phenotyping demonstrated a lambda light chain restricted clonal B-cell population expressing CD20, CD19 and, FMC7. These neoplastic B-cells were negative for CD5 and CD10 expression.

Diagnosis

The morphologic features were consistent with Follicular Lymphoma; however the phenotype (BCL-2 negativity in follicular centers) was unusual for this diagnosis. Fluorescence in situ hybridization (FISH) was negative for an IgH/BCL-2 fusion; however, a BCL-6 rearrangement at the 3q27 locus was detected in 70% of the cells.  Taken together, a diagnosis of Follicular Lymphoma with a BCL-6 rearrangement was given.

Discussion

Follicular lymphoma (FL) is a germinal center derived B-cell neoplasm. The majority of cases exhibit the pathognomonic translocation t(1418)(q32; q21). This translocation leads to overexpression of the anti-apoptotic BCL-2 protein, which can be detected by immunohistochemistry on germinal center B cells. Lymphoma cells are usually positive for germinal center origin markers BCL-6 and CD10 and do not co-express CD5. As exhibited in this case, FL can exhibit biologic heterogeneity and may not express these typical markers. The follicular proliferation with absence of germinal center zonation and tingible body macrophages as seen in this case represents classic morphology of follicular lymphoma but aberrant phenotypic markers [and absence of t(14;18)] may be a pitfall in this diagnosis.

FL with lacking of CD10 expression, BCL-2 expression, and t(14;18) translocation and harboring only BCL-6 positivity with 3q27 rearrangement is rare. Only few such cases have been reported in the literature. Published data reveals that the hallmark t(14;18) translocation is absent in about 10-15% of FL. The majority of these cases are negative for BCL-2 expression, and 9-14% of them demonstrate BCL-6 rearrangement (3q27 locus). While BCL-6 rearrangement can be present in both the usual t(14;18) harboring FL, and also in cases without t(14;18), the latter is rare. Interestingly, studies have shown BCL-6 rearrangements to be more frequent in in BCL-2 rearrangement negative FL – which is evidence of the anti-apoptotic role of non-rearranged BCL-6 in certain microenvironments.

One third of t(14;18) negative FL are also reported to have rare or negative expression of CD10. Morphologically, this subtype has been shown to have significantly larger follicles than  their t(14;18)-positive counterparts, but the distinction may not be obvious in all cases. Some of these cases are shown to have a component of monocytoid B cells. This findings can be problematic in differentiating these FL cases from marginal zone lymphoma (MZL) that can also harbor BCL-6 rearrangements and lack t(14;18), CD10 and BCL-2 positivity. Absence of prominent marginal zone proliferation, BCL-6 protein expression and characteristic genetic alterations present in MZL, such as trisomies 3, 7, and 18 can help differentiating MZL from t(14;18)-negative FL.

This case highlights the importance of morphologic evaluation of a excisional biopsy tissue, and FISH studies to help identify the rare t(14;18) negative FL. While the reported cases are few, there is no published difference in prognosis or survival when compared to t(14;18)-positive FL. As such, it is not clear whether the follicular lymphoma grading scheme applies to t(14;18)-negative FL; however, no significant grading difficulties or differences have been reported.

References

  1. Jardin F, Gaulard P, Buchonnet G, et al. Follicular lymphoma without t(14;18) and with BCL-6 rearrangement: a lymphoma subtype with distinct pathological, molecular and clinical characteristics. Leukemia. 2002;16:2309–2317
    2. Leich E, Salaverria I, Bea S, et al. Follicular lymphomas with and without translocation t(14;18) differ in gene expression profiles and genetic alterations. Blood. 2009;114(4):826-834.

 

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Aadil Ahmed, MD is a 3rd-year anatomic and clinical pathology resident at Loyola University Medical Center. Follow Dr. Ahmed on Twitter @prion87.

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-Kamran M. Mirza, MD PhD is an Assistant Professor of Pathology and Medical Director of Molecular Pathology at Loyola University Medical Center. He was a top 5 honoree in ASCP’s Forty Under 40 2017. Follow Dr. Mirza on twitter @kmirza.

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

Case History

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

Answer

Influenza…but how can that be?

Discussion

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

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

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

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Figure 1. Calculation of positive and negative predictive values.

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

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

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

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

 

References:

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

 

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