How to Detect COVID-19 Variants of Concern

It’s a little deja-vu writing this title one year after a similar blog post on how to validate a COVID-19 assay at the start of the pandemic. In many ways, the challenges are similar: limited reagents/control material, and rising case counts. At least now, there is increasing support in the way of funding from the federal government that could help with monitoring and surveillance. I’m going to summarize the current methods available for detecting the Variants of Concern and emerging variants.

Whole Genome Sequencing

The principle method used by many is whole genome sequencing. It has the advantage of being able to comprehensively examine every letter (nucleotide) of the SARS-CoV-2 genome (30 kilobases long). At our institution, I’ve been working on the effort to sequence all of our positive specimens. While it is achievable, it is not simple nor feasible at most locations. Limitations include:

  • Financial: must already own expensive sequencers
  • Expertise: advanced molecular diagnostics personnel needed who perform NGS testing
  • Data Analytics: bioinformatics personnel needed to create pipelines, analyze data and report it in a digestible format.
  • Timing: the process usually takes a week at best and several weeks if there is a backlog or not enough samples for a sequencing run to be financially viable.
  • Sensitivity: the limit of detection for NGS is 30 CT cycles, which for us includes only about 1/2- 1/3 of all positive COVID19 specimens.

 Bottom line: WGS is the best at detecting new/ emerging strains or mutations when cost/ time is not a concern.

Mutation Screening

Other institutions have begun efforts to screen for variants of concern by detecting characteristic mutations. For instance, the N501Y mutation in the spike protein is common to the major Variants of Concern (UK B.1.1.7, Brazil P.1, and S Africa B.1.351) and E484K is present in the Brazil (P.1), S Africa (B.1.351) and New York Variant (B.1.526). Thus, several institutions (listed below) took approaches to 1) screen for these mutations and then 2) perform WGS sequentially.

InstitutionMethodTargets
Hackensack Meridian Health (HMH)Molecular Beacon Probes, melting tempN501Y, E484K molecular beacons
Rutgers, New JerseyMolecular Beacon Probes, melting tempN501Y molecular beacons
VancouverProbe + melting curve (VirSNiP SARS-CoV-2 Mutation Assays)N501Y screen + qPCR reflex; Probe, melt curve assay
YaleRT-qPCR probe assayS:144del, ORF1Adel
ColumbiaRT-qPCR probe-assayN501Y, E484K

As you can see, HMH, Rutgers and Vancouver are using assays that use probes specific to characteristic alleles combined with melting temperature curves to detect a mutation induced change. Melting curve analysis is normally performed after qPCR to ensure that a single, correct PCR product is formed. This measure is calculated based on the change in fluorescence that occurs when the fluorescent marker is able to bind to its target DNA. Thus the Tm (melting temperature) is similar to the annealing temperature. In this case where a mutation is present in the probe (DNA fragment) binding site, binding is disrupted and occurs at lower a temperature as seen by the downward shift of 5 degrees Celsius in the graph below.

Figure 1. Schematic showing the melting temperature shift for the HMH designed probe binding normal and mutant (E484K variant) sequences at decreasing concentrations.
Figure 2. Similar shift downward in melting temperature for the Rutgers assay when a wild type probe encounters a mutant vs. WT sequence.

These approaches are quick, but can only perform a 2-3 reactions per well and require much of the same expenses as diagnostic RT-qPCR assays. Most of the studies describe this method as a way of screening for samples to be NGS sequenced, however they will not be as good at detecting emerging strains. For example, the N501Y mutation is not present in the New York nor California variants.

Multiplex RT-qPCR can solve some of these problems. At Columbia and Yale, multiple targets are designed to detect B.1.1.7 (N501Y only at Columbia and S144del + ORF1A del at Yale) vs. Brazil/ S. Africa variants (N501Y & E484K at Columbia and ORF1A only at Yale). As new variants have arrived, we found the New York strain carrying both ORF1A deletion and the E484K mutation. It is now clear there are some hotspot areas for mutation within the SARS-CoV-2 genome, which can complicate interpretations. Therefore, these RT-PCR assays are still useful for screening, but do not replace the need for Whole Genome Sequencing.

Genotyping

Given the overlapping spectrum of mutations, it would be helpful to test several markers all at once in a single reaction. At a certain point, this would effectively “genotype” a variant as well as WGS. The assays above have been limited to 2 targets/ reaction due to limited light detection channels. Therefore, I’ve created a multiplex assay that can be scaled up to include 30-40 targets within a single reaction without the need for expensive probes. This method is multiplex PCR fragment analysis, which is traditionally used for forensic fingerprinting or bone marrow transplant tracking. In this method, DNA of different length is amplified by PCR, then separated by capillary electrophoresis-the same instrument that performs Sanger Sequencing.

Fragment analysis can be performed to detect deletion/ insertion mutations and single nucleotide polymorphisms (SNPs) by allele-specific primers or with restriction enzymes that only cut the WT or Mutant sequence.

I designed the assay to target 3 deletion mutations in B.1.1.7: S:D69_70, S: D144, and ORF1A: D3675_3677. Each deletion has a specific length and if 3/3 mutations are present, then there is 95% specificity for the B.1.1.7 strain. Samples from December to present were tested and in the first batch, I detected the characteristic B.1.1.7 pattern (expected pattern and observed pattern below).

Theoretical picture of what the fragment analysis assay would look like for B.1.1.7. An actual patient sample results below, which showed the expected deletions exactly as predicted:

We have tested and sequenced over 500 positive specimens, and we found increasing levels of the B.1.1.7 strain prevalence up to nearly 30% by the middle of March. All screened B.1.1.7 specimens were validated by WGS. These results and the ability to detect the New York and California variants are detailed in our recent pre-print.

Weekly prevalence of isolates consistent with B.1.1.7 in North Texas.

Implications for future Variant Surveillance

As B.1.1.7 has become the dominant strain, and sequencing efforts are increasing. I would argue that assays should be used for what they are best at. For instance, it could be considered a waste of NGS time and resources to sequence all Variants when >50% are going to be B.1.1.7 if other tests can verify the strain faster for 10-20% of the cost. Instead, I think WGS should be focused on discovering emerging variants for which it is best suited. Across the US, case numbers have been decreasing and the number of specimens testable could be expanded by using a more sensitive PCR assay that could.

References

  1. Clark AE et al. Multiplex Fragment Analysis Identifies SARS-CoV-2 Variants. https://www.medrxiv.org/content/10.1101/2021.04.15.21253747v1
  2. Zhao Y et al. A Novel Diagnostic Test to Screen SARS-CoV-2 Variants Containing E484K and N501Y Mutations. A Novel Diagnostic Test to Screen SARS-CoV-2 Variants Containing E484K and N501Y Mutations | medRxiv
  3. Banada P et al. A Simple RT-PCR Melting temperature Assay to Rapidly Screen for Widely Circulating SARS-CoV-2 Variants. A Simple RT-PCR Melting temperature Assay to Rapidly Screen for Widely Circulating SARS-CoV-2 Variants | medRxiv
  4. Annavajhala MK et al. A Novel SARS-CoV-2 Variant of Concern, B.1.526, Identified in New York. A Novel SARS-CoV-2 Variant of Concern, B.1.526, Identified in New York | medRxiv
  5. Matic N et al. Rapid detection of SARS-CoV-2 variants of concern identifying a cluster of B.1.1.28/P.1 variant in British Columbia, Canada. Rapid detection of SARS-CoV-2 variants of concern identifying a cluster of B.1.1.28/P.1 variant in British Columbia, Canada | medRxiv
  6. Vogels CBF et al. PCR assay to enhance global surveillance for SARS-CoV-2 variants of concern. PCR assay to enhance global surveillance for SARS-CoV-2 variants of concern | medRxiv

Jeff SoRelle, MD is Assistant Instructor of Pathology at the University of Texas Southwestern Medical Center in Dallas, TX working in the Next Generation Sequencing lab. His clinical research interests include understanding how lab medicine impacts transgender healthcare and improving genetic variant interpretation. Follow him on Twitter @Jeff_SoRelle.

COVID Variants

Since my last post on the B.1.1.7 (UK) variant, several other variants have arisen. I wanted to describe what makes some Variants of Interest and other Variants of Concern. While a “variant” is often synonymous with a mutation in genetic terms, in the context of SARS-CoV-2, variant means an alternative strain of the virus.

To become a Variant of Interest (VOI), the World Health Organization (WHO) or Centers for Disease Control (CDC) has the following characteristics:

  • Evidence of variants that affect transmission, resistance to vaccines/ therapeutics, mortality, or diagnostic tests
  • Evidence that the variants is contributing to a rise in the proportion of cases in an area.
  • However, limited geographical spread.

Examples: P.2 (from Brazil) B.1.525 (New York), and B.1.526 (New York).

Variants of Concern have increased problems with the same characteristics listed above:

  • Evidence of reduced vaccine protection from severe disease
  • Evidence of substantially reduced response to neutralizing antibodies or therapeutics
  • Evidence of widespread spread
  • Increased Transmissibility or disease severity

Current VOCs: B.1.1.7 (UK), B.1.351 (South Africa), P.1 (Brazil), and B.1.427/ B.1.429 (California).

The initial VOC of B.1.1.7, B.1.351 and P.1 were identified from having increased spread and more mutations than expected, especially in the Spike gene region (Figure 1).

The N501Y mutation in the Spike protein is present in each VOC. It is located at the tip of the protein that binds the ACE2 receptor, increasing binding strength.

So far, vaccines react against the B.1.1.7 variant. However, B.1.351 pseudovirus shows decreased neutralization by both Moderna and Pfizer sera. Specifically, the E484K mutation in the Spike protein confers resistance to neutralizing antibodies. Thus, the strains B.1.351 and P.1 are more likely to be resistant as would any other strain with the E484K variant.

Lastly, the California variant arose as it was found to rise in prevalence from November to February. The key mutations include W152C and L452R, but the significance of this variant is uncertain. However, this variant has begun to spread over much of Southern California and Nevada.

References

  1. Wu K, Werner AP, Moliva JI, Koch M, Choi A, Stewart-Jones GBE, Bennett H, Boyoglu-Barnum S, Shi W, Graham BS, Carfi A, Corbett KS, Seder RA, Edwards DK. mRNA-1273 vaccine induces neutralizing antibodies against spike mutants from global SARS-CoV-2 variants. bioRxiv [Preprint]. 2021 Jan 25:2021.01.25.427948. doi: 10.1101/2021.01.25.427948. PMID: 33501442; PMCID: PMC7836112.
  2. Tada T, Dcosta BM, Samanovic-Golden M, et al. Neutralization of viruses with European, South African, and United States SARS-CoV-2 variant spike proteins by convalescent sera and BNT162b2 mRNA vaccine-elicited antibodies. Preprint. bioRxiv. 2021;2021.02.05.430003. Published 2021 Feb 7. doi:10.1101/2021.02.05.430003
  3. Gangavarapu, Karthik; Alkuzweny, Manar; Cano, Marco; Haag, Emily; Latif, Alaa Abdel; Mullen, Julia L.; Rush, Benjamin; Tsueng, Ginger; Zhou, Jerry; Andersen, Kristian G.; Wu, Chunlei; Su, Andrew I.; Hughes, Laura D. outbreak.info. Available online: https://outbreak.info/ (2020)
  4. https://www.cdc.gov/coronavirus/2019-ncov/cases-updates/variant-surveillance/variant-info.html

Jeff SoRelle, MD is Assistant Instructor of Pathology at the University of Texas Southwestern Medical Center in Dallas, TX working in the Next Generation Sequencing lab. His clinical research interests include understanding how lab medicine impacts transgender healthcare and improving genetic variant interpretation. Follow him on Twitter @Jeff_SoRelle.

Will the B.1.1.7 variant evade the Vaccine/Tests?

Will the B.1.1.7 variant evade the vaccine/tests?

This question came up recently and I wanted to share some cutting edge information the addresses this. This was in part adapted from Akiko Iwasaki’s (Yale HHMI immunologist) Twitter discussion of this subject.1

Will B.1.1.7 evade our tests?

The UK variant commonly called lineage B.1.1.7 (officially Variant of Concern 202012/01) has 23 genetic variants that result in 17 protein coding changes.2 Most tests including the ones at our institution (Abbott) are not currently affected (see below). Only the ThermoFisher assay has declared a target that covers the 69-70del variant in the S gene (in green). This conversely makes the TaqPath® assay one way to detect a potential B.1.1.7 variant.

Figure 1. A picture of the SARS-CoV-2 genome with red lines indicating mutation sites and different assays and relative location of their qPCR targets.

Will the vaccine protect against the B.1.1.7 variant?

The Pfizer and Moderna RNA vaccines create an immune response against the spike protein. We don’t know the exact sequences or reactivity of the vaccines’ spike protein. However, a recent study looked at the antibody reactivity to linear epitopes of COVID-19 in 579 patients who were naturally infected with COVID-19. For the antibodies against the spike, the major reactive linear epitopes are indicated in Red at the bottom. None of the B.1.1.7 mutations (Orange) overlap with these major reactive epitopes.3 

Figure taken from Reference 3.

For a closer look, see below.

Figure taken from Reference 3.

A limitation of these analyses is the use of only linear epitopes. Mutations might impact a 3D epitope affecting Ab binding. However, people make multiple antibodies to the spike protein.4 So, broad coverage should arise after exposure to the either the vaccine or natural infection with COVID-19.

The vaccine should induce a polyclonal antibody response that recognizes multiple parts of the spike protein, making it effective, even against novel variants. Also, there should be few to no False Negative COVID-19 tests due to the new variant, but we will continue to monitor and test this experimentally. 

References

  1. Prof. Akiko Iwasaki @VirusesImmunity
  2.  Chand, Meera et al. Investigation of novel SARS-COV-2 variant: Variant of Concern 202012/01 Public Health England.
  3. Haynes WA et al. High-resolution mapping and characterization of epitopes in COVID-19 patients. MedRxiv. https://www.medrxiv.org/content/10.1101/2020.11.23.20235002v1#p-5
  4. Shrock E et al. Viral epitope profiling of COVID-19 patients reveals cross-reactivity and correlates of severity. Science 2020 370(6520). https://science.sciencemag.org/content/370/6520/eabd4250

Jeff SoRelle, MD is Assistant Instructor of Pathology at the University of Texas Southwestern Medical Center in Dallas, TX working in the Next Generation Sequencing lab. His clinical research interests include understanding how lab medicine impacts transgender healthcare and improving genetic variant interpretation. Follow him on Twitter @Jeff_SoRelle.

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

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

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

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

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

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

Here are 3 suggestions:

1) Establish testing algorithms and clear sample workflows.

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

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

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

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

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

2) Communicate to clinical staff frequently about your tests.

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

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

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

3) Keep the lab staff off the phone.

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

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

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

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

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

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

Monitoring Bone Marrow Transplant Recipients

Hello everyone, it’s been quite some time since my last post. I hope everyone has remained safe and healthy during these times!

My last post dived into short tandem repeat (STR) analysis for bone marrow engraftment monitoring. Today is a presentation of a patient who was transplanted for treatment of acute myeloid leukemia (AML). With all patients (with minor exceptions), donor and pre-transplant recipient samples are taken before transplant. Their informative alleles are then identified and used to determine the percent of donor and any recipient cells in subsequent post-transplant samples.

This patient was unique in that we were not able to obtain the donor sample (they were transplanted outside of our system), and therefore we used a buccal swab for their pre-transplant recipient informatives.

Buccal swabs are chosen because they are a non-invasive way to obtain squamous epithelial cells. These cells are important because they are of the recipient origin and will not change. With this technique, it is essential that the patient has no mucosal inflammation or is not too rough when swabbing their cheek. Otherwise, the buccal sample may become contaminated with blood which would contain donor cells.

We then inferred the donor informatives from the data of a mixed sample and the buccal swab.

Calculation of recipient and donor percentage in a post-transplant sample is determined on specific formulas that utilize these informative alleles. But what happens when a patient relapses and new mutations or deletions are introduced into their genome, causing a change in these informative alleles?

In this case, the patient had a loss of allele at two loci (CSF1PO – allele 11 and D5S818 – allele 13) after having previously obtained full engraftment (Figure 1).

Figure 1. The pre-sample was acquired through a buccal swab. There was no donor sample that was acquirable, and therefore the donor informative alleles were inferred through available data. In September of 2019, the patient was at 100% donor. Almost a year later, the patient is now at 4% donor and missing previously identified recipient alleles, indicating a loss of allele/mutation. Brown box with “R” stands for recipient. Blue box with “D1” stands for donor. Green box with D1R stands for shared.

The importance here is that the true percent donor is 4% (Figure 2). If we take a look at the affected informative alleles, we see an erroneous result of 100% donor and NI (which means the locus is non-informative, eliminating it from the calculations). This expands on the importance of an analyst to be attentive to the results presented. While this case was clearly evident and was caught by our error measurements, it is theoretically possible to cause an issue, especially in cases where the recipient percentages may be smaller. Furthermore, this phenomenon stresses the importance of including multiple informative alleles in our analysis, which increases our measurement of confidence.1

Figure 2. CSF1PO and D5S818 are incorrectly representing the patient’s status. CSF1PO is representing the patient at 100% donor and D5S818 is automatically identified as a non-informative by our software. After automatic and manual loci ignores, the total percent donor was 4%

We know that a loss of allele (loss of heterozygosity) is the likely explanation because both loci are in locations specific to the disease. Looking at Figure 3 below, the two alleles were affected because they were both present on the long arm of chromosome 5. Further, this chromosome is known to be involved in AML, and is also, of course, associated with other disorders like MDS.2 Additionally, the patient had cytology testing that identified this as an affected chromosome.

Figure 3. CSF1PO and D5S818 are both located on the long arm of chromosome 5. CSF1PO’s location is 5q33.1 and D5S818’s location is 5q23.2.

This is an interesting phenomenon and one that shows in measurable terms how a patient’s status can affect their molecular results. It’s further an expression of the molecular mechanisms of a disease, one of my first measurable experiences of how a disease affects the physical molecular constituents of another human.

To me, this encounter was an expression of how complicated, and yet connected, the entire genome has been designed. I am continuously amazed and look forward to expanding my understanding of molecular science.

References

  1. Crow J, Youens K, Michalowski S, et al. Donor cell leukemia in umbilical cord blood transplant patients: a case study and literature review highlighting the importance of molecular engraftment analysis. J Mol Diagn. 2010;12(4):530-537. doi:10.2353/jmoldx.2010.090215
  2. Crow J, Youens K, Michalowski S, et al. Donor cell leukemia in umbilical cord blood transplant patients: a case study and literature review highlighting the importance of molecular engraftment analysis. J Mol Diagn. 2010;12(4):530-537. doi:10.2353/jmoldx.2010.090215

-Ben Dahlstrom is a recent graduate of the NorthShore University HealthSystem MLS program. He currently works as a molecular technologist for Northwestern University in their transplant lab, performing HLA typing on bone marrow and solid organ transplants. His interests include microbiology, molecular, immunology, and blood bank.

Massive COVID-19 Testing: 30 Million Tests/Week

Population COVID-19 testing

Population-wide testing to identify symptomatic and asymptomatic infections could be a powerful tool to control Coronvirus Disease 2019 (COVID-19) spread, but current global testing capacity does not permit widespread testing of asymptomatic individuals. These tests are still limited to individuals who are symptomatic with limited availability to those with recent exposure to an infected person.

Because of the high prevalence of asymptomatic COVID-19 infections, proposals from the Rockefeller Foundation for disease mitigation include widespread and frequent testing of the US population. In the United States, diagnostic testing for SARS-CoV-2, the causative virus of COVID-19 is currently >2 million per week. Estimates for US testing needs for population wide surveillance range from 30 to 300 million per week. In order to scale testing by an order of magnitude, novel technologies and rethinking current testing paradigms are needed. The NIH has initiated a rapid funding program to develop SARS-CoV-2 testing, and these new technologies may play a part. However, we can broadly conceptualize key problems to address in population-wide testing in the US. The first is high-sensitivity testing which identifies active infection and can be performed with massive throughput. The second is the logistics of gathering hundreds of thousands of samples to each testing laboratory each day.

Next Generation Solutions to COVID testing

Emerging technologies using targeted next-generation sequencing have been suggested as a potential solution to population-wide testing. The key features include 1) extraction free amplification 2) an easily collected specimen such as saliva, 3) nucleotide barcodes to enable sample pooling, and 4) a limited number of targets (to allow deeper sequencing, i.e. higher sensitivity). Illumina is selling a whole genome test for SARS-CoV-2, but this limits sequencing to 3,000 tests/ run. Another recent approval for a private testing lab uses only one target, and may allow it to increase to 100,000 tests/ day. And a recent protocol for LAMP-Seq in pre-print outlines how this could work in a scheme below. An attractive aspect of this approach is decentralized specimen processing.

Whereas Bill Gates has supported a portfolio approach to vaccines placing multiple bets on different processes in parallel, a similar approach should be applied to multiplexed sequencing methods. Two sequencing runs can be performed on a single instrument in a single day, which can process several thousand samples. However, sequencing is not the only step in sequencing; library preparation and specimen handling take significant amounts of time too.

Laboratory Logistics

This technology would represent an exponential expansion in analytic testing capacity, but clinical labs will require a similar escalation in logistic capacity. The largest clinical laboratories in the world process less than 100,000 samples per day. Clinical laboratories have a long history of automation with the first robotic specimen track systems developed in the 1980s. Engineering and clinical lab expertise should thus partner to innovate on methods to handle high volumes. This level of investment for an issue that is likely to fade in 2 years, is not attractive to most private health systems, so public investment from multiple states in regional reference labs is needed.

It is still hard to conceive the necessary scale up in sample processing can be achieve within the time frame needed, so I would also propose a de-centralized sample processing approach. This would include self-collection of saliva (a safe, effective sample type with similar sensitivity as nasopharyngeal swabs), drop-off sites, and processing at places like Pharmacies (>90% of Americans live within 5 miles of a pharmacy and they could be authorized to administer tests- just as they administer vaccines). This would introduce pre-analytic problems, but if the goal is frequent and high rates of testing, then we will have to accept certain losses in sensitivity (which currently is arguably better than it needs to be). Interestingly, pre-analytic concerns with saliva have not led to sample instability or degradation of RNA causing false negatives, as described in my last post. However, other factors could affect saliva quality: smoking, age, and genetic factors of water: protein ratio affecting viscosity.

Testing solutions should be considered in the context of the planned testing network. The specimen type should be easy for the patient to provide, processed with existing laboratory equipment and resulted electronically. For example, current COVID-19 testing is based on sample collections requiring a healthcare worker encased in personal protective equipment (PPE) utilizing a swab device. Testing needs to progress to a simpler solution such as saliva which can be collected by the patient in the absence of a swab or PPE. Preliminary studies have demonstrated that saliva is sample type comparable to nasopharyngeal swab. The ideal saliva sample would be collected into an existing collection tube type (e.g. red-top tubes) which are already compatible with existing laboratory automation. In aggregate, a person could spit into a tube at-home, have the tube sent to a laboratory, and in the laboratory the tube would be directly placed onto an automated robotic track system. 

Laboratory professionals need to provide a comprehensive plan for regional and national laboratory networks which can scale to provide overwhelming force to COVID-19 testing. No other profession or governmental organization understands testing as much as we do. Our understanding of managing samples from collection to result should be applied to the pandemic at hand. Until now most laboratorians in the US have focused on the immediate needs of providing testing for symptomatic patients and healthcare workers.

Vision for automated COVID-19 testing

One could envision an automated line of testing that moves samples through processing to allow multiplexing and combinations of samples to allow large numbers of patients to be tested at once (see below). This is feasible in some specialized centers, but would require investments in automation, bioinformatics, and interfaces for a seamless process (figure below). If testing mostly asymptomatic patients, it may also be possible to do this on pooled samples. The number of samples to pool would depend on the likelihood to having a positive result (this would require sequencing all individuals in a pool).

This represents a synthesis of ideas in decentralized specimen collection, laboratory automation and massive testing throughput with Next-Generation Sequencing, but unfortunately this is not yet a reality.

References

  1. Jonathan L. Schmid-Burgk et al. LAMP-Seq: Population-Sclae COVID-19 Diagnostics Using Combinatorial Barcoding. bioRxiv 2020.04.06.025635.
  2. The Rockefeller Foundation. National Covid-19 Testing Action Plan Pragmatic steps to reopen our workplaces and our communities. 2020.
  3. Cahill TJ, Cravatt B, Goldman LR, Iwasaki A, Kemp RS, Lin MZ et al. Scientists to Stop COVID-19.  OR Rob Copeland, Wall Street Journal (2020) The Secret Group of Scientists and Billionaires Pushing a Manhattan Project for Covid-19. April 27
  4. https://www.illumina.com/products/by-type/ivd-products/covidseq.html

-Jeff SoRelle, MD is a Chief Resident of Pathology at the University of Texas Southwestern Medical Center in Dallas, TX. His clinical research interests include understanding how the lab intersects with transgender healthcare and improving genetic variant interpretation.

False Negatives in COVID-19 Testing

I left for vacation at the beginning of June thinking “once I get back, all of this COVID stuff will be quieted down.” …Well that wasn’t quite the case and testing for novel Coronavirus has continued to be very important. In fact, this last weekend I was tested by occupational health. It came back negative, but I’m am very enthusiastic to get alternative specimen types validated; those Nasopharyngeal swabs are quite…uncomfortable. Luckily, my test was processed at our institution which gets results back in 24-48 hours. However, with the resurgence around the country, turnaround times are backing up to 7-8 days. One solution has been the widely used IDNOW point of care platform. However, there has been significant concern over false negatives produced by this platform. One reason the sensitivity is different is because this platform performs isothermal amplification of nucleic acid. This method amplifies RNA at a stable temperature instead of cycling the temperature as in real-time PCR.

Colleagues at my institution reflexed any negative IDNOW samples to the m2000 Real-Time PCR assay for SARS-CoV-2 for one month. Within that time, over 500 samples were tested and the IDNOW was found to have missed 21% of positive cases (prevalence rate of 5%)2. One the positive side, it had a 98% negative predictive value, which helped rule out COVID19 infection. However, as prevalence rates are increasing, a high negative predictive value isn’t as important as sensitivity.

One study drew much attention when it claimed the IDNOW had a sensitivity of 52% in a New York City academic institution (Basu)4. However, this seems to be an outlier compared to other studies of this platform: one large multi-center study found positive percent agreement (equivalent of sensitivity when a gold standard test hasn’t been established) of 74%1. The highest PPA of 88%3 for the IDNOW was found in a study that indicated it can be completed in 17 minutes, whereas another quick instrument (but not point of care instrument: Xpert Xpress, 45min) had a PPA of 98%2.

Myself and other colleagues looked more closely at the clinical characteristics of false negative test results on the IDNOW. Overall, we found 82% PPA, and 8 patients with false negative tests. Interestingly, a majority of these patients were tested over 2 weeks after their initial onset of symptoms. The virus is known to be at its highest levels at the beginning of symptom onset. So the test may not be limited, but it should be used in the correct clinical context (< 2weeks from symptom onset). After that time, other RT-PCR based tests are more appropriate.

As clinical laboratorians, we often hear: “the right test for the right patient at the right time.” Now with so many platforms available for use in different contexts, we should help guide clinicians to Choose Wisely.

References

  1. Harrington A et al. Comparison of Abbott ID Now and Abbott m2000 methods for the detection of SARS-CoV-2 from nasopharyngeal and nasal swabs from symptomatic patients. JCM 2020. PMID: PMID: 32327448
  2. McDonald et al. Diagnostic Performance of a Rapid Point of Care Test for SARS-CoV-2 in an Urban ED Setting. Academ. Emerg. Med. 2020. PMID: 32492760
  3. Zhen W et al. Clinical Evaluation of Three Sample-To-Answer Platforms for the Detection of SARS-CoV-2. JCM 2020. PMID: 32332061
  4. Basu A et al. Performance of the rapid Nucleic Acid Amplification by Abbott ID NOW COVID-19 in nasopharyngeal swabs transported in viral media and dry nasal swabs, in a New York City academic institution. BioRxiv 2020.

-Jeff SoRelle, MD is a Chief Resident of Pathology at the University of Texas Southwestern Medical Center in Dallas, TX. His clinical research interests include understanding how the lab intersects with transgender healthcare and improving genetic variant interpretation.

Biomarker Testing for Cancer Patients: Barriers and Solutions Part 6

This month we will finish the discussion the common barriers to biomarker testing for cancer patients in the community. Lengthy complex reports is a relatively straightforward barrier to address, so I will pair it with the lack of education on guidelines barrier to complete this blog series on barriers to biomarker testing.

As you may recall, these are the top 10 barriers that I’ve seen to biomarker testing in the community:

  1. High cost of testing.
  2. Long turnaround time for results.
  3. Limited tissue quantity.
  4. Preanalytical issues with tissue.
  5. Low biomarker testing rates.
  6. Lack of standardization in biomarker testing.
  7. Siloed disciplines.
  8. Low reimbursement.
  9. Lengthy complex reports.
  10. Lack of education on guidelines.

Lengthy Complex Reports

Laboratory issued reports are typically developed by the lab and are often written in a manner that is easy to understand for other laboratorians. I’m guilty of writing long interpretive comments that are attached to every molecular diagnostics results. I would get irritated when the physician would call and ask questions that in my mind were clearly addressed in the interpretive comment. I thought the issue was they were not reading the comments (and this could be true). I now understand that the issue is that the comments were not written for the end-user.

When insourcing NGS I was fortunate enough to get feedback from the multidisciplinary team in the Molecular Steering Committee. One of the complaints that I heard loudly locally, that also resonated in the community, was the reports for NGS were way too long and they didn’t find value in half of the information that was in the report. When were shopping for the right cloud-based reporting software, I kept the feedback in mind from the oncologists. I was actually able to get proto-type reports from 3 different companies and provide them to the oncologists for them to score and provide feedback on the layout. This was invaluable in developing a report that worked well for the treating physician and not the laboratory.

Some of the feedback they gave that made a direct impact into the report we created was: bold the patient’s name so they can easily find it, use patterns as well as color-coding for drug resistance/sensitivity in case the document is faxed, and tell them everything they need to know to make treatment decisions on page one. These are things that were not intuitive to me. Having end-user feedback helped us generate a more useable report and enlightened me that the report needs to be written to an oncology audience.

Lack of education on guidelines

I’ve had the opportunity to do a great deal of educating around biomarker testing in the community. Physicians and nurses in the community want to provide guideline-driven care. Often when we are educating on changes to guidelines, it’s the first time the providers have heard of the change. NCCN for lung cancer alone had at least 7 updates in 2019. It’s amazing that the guidelines are able to keep up with the ever changing science and drug approvals; however it’s incredibly difficult to keep track of the changes.  

In large institutions we are fortunate enough to have specialized physicians that help keep the rest of us informed of changes in their area of expertise. Community physicians typically see and treat all types of cancers and don’t always have the network of specialists to keep them informed of changes for every cancer type. Many of them also do not have the time to attend conferences due to heavy workload.

 In order for the community physician to be informed of all of the changes to guidelines for every tumor type, we need to make sure the information is provided in a variety of methods. The information needs to be easily accessible. I have found that educational programs work well when brought to the community rather than trying to get the community to come to them. Pharmaceutical and diagnostic companies and even reference laboratories now have teams of individuals in roles that are intended to educate and not sell. They can provide in office education, facilitate webinars, lunch and learns, and dinner programs. If there is a champion for biomarker testing within the facility, you can develop your own educational program to be delivery locally at grand rounds. We discuss changes to guidelines within our Molecular Steering Committee. I’ve also talked to institutions where this education is given during tumor boards.

I don’t think there is a bad forum for education. Some physicians may prefer getting guideline updates from twitter; others will be more comfortable with a discussion with an expert, regardless of the medium it is important that we help facilitate education of guidelines in order to increase biomarker testing rates in the community.

-Tabetha Sundin, PhD, HCLD (ABB), MB (ASCP)CM,  has over 10 years of laboratory experience in clinical molecular diagnostics including oncology, genetics, and infectious diseases. She is the Scientific Director of Molecular Diagnostics and Serology at Sentara Healthcare. Dr. Sundin holds appointments as Adjunct Associate Professor at Old Dominion University and Assistant Professor at Eastern Virginia Medical School and is involved with numerous efforts to support the molecular diagnostics field. 

Extraction-free and Saliva COVID-19 Testing

Much has changed quickly with SARS-CoV-2 virus (COVID-19) testing. Several commercial options are now available. Labs have less problems getting control material (positive samples are no longer in short supply). And labs that opted to bring on testing are now running multiple versions of COVID-19 molecular tests with a combination of high speed platforms or high throughput. Rapid cartridge tests are used for clearing people from the ED/ removing contact isolation on inpatients while the high throughput assays are used for routine screening.

However, several bottlenecks still exist. There are shortages of nucleic acid extraction kits, collection swabs and viral transport media. Fortunately, some recent studies have demonstrated preliminary evidence for using alternative sample types, collection methods, and storage conditions.

One of the first tenets of molecular diagnostics is isolation and purification of nucleic acid. Therefore, it was surprising to see a report on an extraction-free COVID-19 protocol from Vermont (Bruce EA et al.). This study initially analyzed two patient samples and showed drops in sensitivity of ~4Ct cycles. While this would not be suitable for low level detection, many viral samples have high levels of virus that still would permit detection. The team went on to test this method on 150 positive specimens from the University of Washington and found 92% sensitivity with 35% sensitivity at the low viral load range (Ct value> 30). This was improved with a brief heat inactivation step (Table 1). This was similarly seen in a study from Denmark, where brief heat inactivation of extraction-free methods (Direct) had 97% specificity in 87 specimens (Table 2).

Table 1. Vermont study comparing sensitivity of direct RT-PCR (no extraction step) with the validated results of 150 specimens coming from the University of Washington.
Table 2. Denmark study found extraction-free protocols (Direct) were comparable to extracted RNA (MagNA Pure extraction method) detection in 87 specimens.

Some similar studies out of Chile also showed extraction-free protocols on a larger number of specimens, and they reported a loss in sensitivity varying from 1-7 Ct cycles depending on the primers used.

Figure 1. P1 and P2 are patient 1 and 2. NSS indicates a nasal swab sample where RNA was extracted. RNA indicates a sample with no RNA extraction.

As this novel Coronavirus has an RNA-based genome, RNA is the target of molecular tests. As RNA is susceptible to degradation, there have been concerns over sample storage. Should it be refrigerated? Frozen? How do multiple freeze-thaw cycles impact specimen stability? Are there viable alternatives to viral transport media? One preliminary study explored these questions very nicely. They took X multiple sample types (NP, BAL, saline storage media) and stored them at 20C, 4C, -20, and -70 for multiple days up to 1 week and then analyzed the level of virus detected. In each case, the loss in sensitivity was minimal (<2 Ct cycles from day 0 to day 7) at room temperature with comparable results at lower temperatures (Table 3).

Table 3. Stability of SARS-CoV-2 RNA detected by the Quest EUA rRT-PCR. VCM- viral culture media; UTM-R Copan’s transport medium; M4-microtest media; BAL- bronchoalveolar lavage.

Lastly, alternative sample types such as saliva will help break the bottleneck in swabs and viral transport media. I was surprised to hear about this being a suitable alternative. Having worked with saliva for DNA analysis, I know it can be contaminated, of variable quantity, includes digestive enzymes and is viscous (slimy). These are not characteristics a lab would look for in a specimen type being used for high-throughput testing where several sample failures could occur. But these researchers from Yale showed measurable levels of SARS-CoV-2 that facilitated even higher sensitivity than nasopharyngeal swabs (Wylie AL et al).

Figure 2. SARS-CoV-2 titers are higher in the saliva than nasopharyngeal swabs from hospital inpatients. (a) All positive nasopharyngeal swabs (n = 46) and saliva samples (n = 39) were compared by a Mann-Whitney test (p < 0.05). Bars represent the median and 95% CI. Our assay detection limits for SARS-CoV-2 using the US CDC “N1” assay is at cycle threshold 38, which corresponds to 5,610 virus copies/mL of sample (shown as dotted line and grey area). (b) Patient matched samples (n = 38), represented by the connecting lines, were compared by a Wilcoxon test (p < 0.05). (c) Patient matched samples (n = 38) are also represented on a scatter plot.

With a much-needed increase in testing for this country, optimizations need to be implemented to improve efficiency. These steps alone will not be enough, but if we can have extraction-free testing of saliva collected at home, this would provide a substantial benefit to bringing easy testing to everyone.

UPDATE: Since this was written, the first FDA EUA was authorized for an at-home saliva collection kit for use at the Rutger’s clinical genomics lab (https://www.fda.gov/media/137773/download).

References

Please note: many of these references were on pre-print servers and have not been peer-reviewed.

  1. Bruce EA, Huang ML, Perchetti GA, et al. DIRECT RT-qPCR DETECTION OF SARS-CoV-2 RNA FROM PATIENT NASOPHARYNGEAL SWABS WITHOUT AN RNA EXTRACTION STEP. 2020. https://www.biorxiv.org/content/10.1101/2020.03.20.001008v2.full#T2
  2. Wyllie AL, Fournier J, Casanovas-Massana A, Campbell M et al. Saliva is more sensitive for SARS-CoV-2 detection in COVID-19 patients than nasopharyngeal swabs. medRxiv 2020. https://www.medrxiv.org/content/10.1101/2020.04.16.20067835v1#disqus_thread
  3. Fomsgaard AS, Rosentierne MW. An alternative workflow for molecular detection of SARS-CoV-2 – escape from the NA extraction kit-shortage, Copenhagen, Denmark, March 2020. https://www.medrxiv.org/content/10.1101/2020.03.27.20044495v1.full.pdf
  4. Rogers AA, Baumann RE, Borillo GA, et al. Evaluation of Transport Media and Specimen Transport Conditions for the Detection of SARS-CoV-2 2 Using Real Time Reverse Transcription PCR. JCM 2020.
  5. Beltran-Pavez C, Marquez CL, Munoz G et al. SARS-CoV-2 detection from nasopharyngeal swab samples without RNA extraction. bioRxiv 2020. https://www.biorxiv.org/content/10.1101/2020.03.28.013508v1.full.pdf

-Jeff SoRelle, MD is a Chief Resident of Pathology at the University of Texas Southwestern Medical Center in Dallas, TX. His clinical research interests include understanding how the lab intersects with transgender healthcare and improving genetic variant interpretation.

Introduction to Short Tandem Repeat

Hello again!

I hope everyone is staying safe and healthy during these unfortunate times. My last post was in relation to being a new MLS grad and beginning my career as a molecular technologist at Northwestern University’s Transplant Lab. Time definitely flies by!

Today I’m going to provide a basic introduction on an assay I’ve recently been trained on called Short Tandem Repeat (STR). If you were to take a glance at your genome, it would be littered with many repeating sequences. While there are many different classifications of repeating sequences, STRs are a type of tandem repeating sequence where each repeat is approximately 2 to 7 nucleotides in length.1,2,3 STR is well-known in forensic science to help identify a suspect at a crime scene when different sources of DNA are present. Yet, its applications are many – from cell line confirmation, paternal testing, and all the way to chimerism analysis!4

Image 1. Electropherogram depicting two different alleles (11 and 17) within 1 locus (D6S1043). Allele 11 has 11 repeats and allele 17 has 17 repeats.

STRs are polymorphic, one useful characteristic among many, which make its utilization in identifying the source of DNA particularly advantageous. An STR allele is defined by the number of times the repeating sequence, defined above, repeats. (Image 1).1 Individuals are either heterozygous or homozygous at each locus. As the number of STR loci being evaluated increases, the statistical power of discrimination increases and the likelihood of another individual having the same profile becomes increasingly unlikely and detecting small differences increases.3,4 In our lab, we evaluate a total of 21 different loci!

Additionally, in our HLA lab we utilize STR to monitor chimerism status in patients who have undergone an allogeneic stem cell transplant. Before their transplant, patients are matched to a donor through their HLA system (different from STR). Once an HLA match is confirmed, we utilize the patient’s pre-transplant and the donor’s sample to generate STR informative alleles. Informative alleles are alleles that are present only in the recipient and not the donor. These alleles are important because stem cell transplants replace the recipient’s marrow and the detection of recipient DNA in post-transplant samples is crucial to identifying rejection or relapse of their disease. Additionally, loci that contain informative alleles are defined as informative loci, these are the loci then used to identify the percentage chimerism (Image 2).

Image 2. Donor and recipient (pre-transplant) are represented in the first two electropherograms. The green “D1R” tags represent shared alleles, the blue “D1” tags represent donor specific alleles, and the brown “R” tags represent recipient specific alleles. In the first locus, AMEL, there are no informative alleles and therefore it is not an informative locus. In the next locus, D3S1358, there is one shared allele, 15, one donor allele, 17, and one recipient informative allele, 18. Informative loci have recipient informative alleles and can detect the presence of recipient DNA in a sample – in this case all of the following loci after AMEL are informative loci. CD3 (post-transplant) is represented on the third electropherogram. In this example, it is clear that the patient is having some sort of graft failure or reoccurrence of their disease because their own cells, instead of just the donor, are present.

When recipient cells begin to re-emerge, we can detect the relative allele peaks and assign them to the recipient or donor by referring to the informative alleles. Allele peaks are then utilized through equations in our software that measure the area under these defined peaks and then compute a donor percentage chimerism. Once that informative report is created, we can compare any proceeding post-transplant sample to determine the patient’s chimerism status (Image 2).

Interesting enough, we don’t simply isolate the DNA from the post-sample buffy like we would with other samples. Rather, we separate each post sample into a total of three sub-samples. The first is simply the patient’s peripheral blood – nothing fancy. The other two are isolated from the rest of the peripheral blood that was not used into two separate cell lineages – lymphocytes (CD3+) and myelocytes (CD33+). This process is extremely labor-intensive, being able to process a set of up to 8-12 patients at a time and each set taking up to 4-5 hours.

The process begins by aliquoting peripheral blood for DNA isolation (sample 1) and taking the remainder and layering it over a lymphocyte separation medium (LSM). Then harvesting the lymphocyte/white cell layer from the spun down LSM. We then go on to add CD3 and CD33 antibody selection cocktails and magnetic beads. Then, we do a series of washes with magnets and eventually end up with our purified CD3 and CD33 cell populations. Their purity is determined through flow cytometry, an important component to confirm that our leukocyte subsets aren’t contaminated with other leukocyte populations – as contamination would defeat the purpose of analyzing different lineages.

Finally, we take the isolated DNA from the three sub-samples and amplify it with specific primers and fluorescent tags through PCR. Then the samples are loaded onto a capillary electrophoresis instrument. This instrument will detect each fragment length, defined by the primers and the repeats within, and be able to identify these fragments through size, fluorescent tags, lasers, and detectors. The instrument will then generate data that we can take to our analyzing platform, which is ChimerMarker. Through this we can analyze the data and generate our clinical reports.

Though extremely time intensive, lineage-specific chimerism is critical in stem cell transplant because it is more informative and sensitive than total leukocyte analysis – being several magnitudes more sensitive than analyzing just peripheral blood alone. It permits early detection of small chimeric cell populations that otherwise may go undetected, as one subset in the peripheral blood may “mask” another subset that has increasing percent recipient cells. Diagnosing these small cell chimeric cell populations as early as possible is critical for therapeutic interventions and reductions in graft rejections.2,5,6

Furthermore, not only is their detection important, but through our analysis we can calculate the percentage of donor cells and recipient cells. We oftentimes report out the donor percentage (%) chimerism. For example, a patient at 322 days post-transplant could have a donor chimerism of 96% in their peripheral blood, 100% in their CD33 lineage, and 73% in their CD3 lineage. Then, at day 364 post-transplant they may then be at 100% in their peripheral blood, 100% in their CD33 lineage, and 92 percent in their CD3 lineage. Two things to notice in this example is that the percentages are changing (increasing in donor chimerism in this case) and that the peripheral blood expressed 100% chimerism in the second sample at 364 days, but when we look specifically at the CD3 sub-population at 364 days there was still 8% of recipient cells present (Image 3 & 4).

Image 3. Samples at 322 days post-transplant. Peripheral blood reports 96% donor chimerism. CD3 and CD33, purified from peripheral blood, reports 73% and 100% donor chimerism, respectively.
Image 4. Samples at 364 days post-transplant, same patient as in Image 3 above. Peripheral blood reports 100% donor chimerism. CD3 and CD33, purified from peripheral blood, reports 92% and 100% donor chimerism, respectively.

Some studies have focused not only on the trends in percentages changing, but also in their relative percentage constellation. For example, one study found that increased recipient CD3 cells had an increased predictive factor of graft rejection. It was also found that further sub-leukocyte populations increased this predictive power.5 Even more, there have been some studies that have looked at chimerism and its usefulness in predicting graft versus host disease (GvHD). This disease is defined by donor leukocytes attacking the leukocytes and tissues of the recipient. Through these and other findings, the potential and applicability of chimerism monitoring is extremely crucial to patient care during their transplant progression.2,5,6,7

While engraftment is a very dynamic process, varying from individuals and disease-types, engraftment monitoring is one way to monitor and ultimately influence therapeutic approaches.2,5,6 I am proud to be able to contribute to the wonderful team here at Northwestern University and I strive to learn more about the process – both clinical and in the lab. In future articles, I hope to go into more detail about the process and other assays that we perform.

Thanks for reading! Until next time! Stay well and safe during these uncertain times!

References

  1. Life Technologies. 2014. DNA Fragment Analysis by Capillary Electrophoresis. Thermo Fisher Scientific. http://www.thermofisher.com/content/dam/LifeTech/global/Forms/PDF/fragment-analysis-chemistry-guide.pdf.
  2. Kristt, D., Stein, J., Yaniv, I., & Klein, T. (2007). Assessing quantitative chimerism longitudinally: technical considerations, clinical applications and routine feasibility. Bone Marrow Transplantation, 39(5), 255–268. doi: 10.1038/sj.bmt.1705576
  3. Clark, J.R., Scott, S.D., Jack, A.L., Lee, H., Mason, J., Carter, G.I., Pearce, L., Jackson, T., Clouston, H., Sproul, A., Keen, L., Molloy, K., Folarin, N., Whitby, L., Snowden, J.A., Reilly, J.T. and Barnett, D. (2015), Monitoring of chimerism following allogeneic haematopoietic stem cell transplantation (HSCT): Technical recommendations for the use of Short Tandem Repeat (STR) based techniques, on behalf of the United Kingdom National External Quality Assessment Service for Leucocyte Immunophenotyping Chimerism Working Group. Br J Haematol, 168: 26-37. doi:10.1111/bjh.13073
  4. Short Tandem Repeat Analysis in the Research Laboratory. (2012). Retrieved April 10, 2020, from https://www.promega.com/resources/pubhub/short-tandem-repeat-analysis-in-the-research-laboratory/
  5. Breuer, S., Preuner, S., Fritsch, G., Daxberger, H., Koenig, M., Poetschger, U., … Matthes-Martin, S. (2011). Early recipient chimerism testing in the T- and NK-cell lineages for risk assessment of graft rejection in pediatric patients undergoing allogeneic stem cell transplantation. Leukemia26(3), 509–519. doi: 10.1038/leu.2011.244
  6. Buckingham, L. (2012). Molecular diagnostics: fundamentals, methods, and clinical applications. Philadelphia: F.A. Davis Company.
  7. Rupa-Matysek, J., Lewandowski, K., Nowak, W., Sawiński, K., Gil, L., & Komarnicki, M. (2011). Correlation Between the Kinetics of CD3 Chimerism and the Incidence of Graft-Versus-Host Disease in Patients Undergoing Allogeneic Hematopoietic Stem Cell Transplantation. Transplantation Proceedings43(5), 1915–1923. doi: 10.1016/j.transproceed.2011.02.011

-Ben Dahlstrom is a recent graduate of the NorthShore University HealthSystem MLS program. He currently works as a molecular technologist for Northwestern University in their transplant lab, performing HLA typing on bone marrow and solid organ transplants. His interests include microbiology, molecular, immunology, and blood bank.