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Published: July 2010Print Record of Viewing
In Part 1, Dr. Binnicker provides an overview and case studies of the conventional methods for detection and diagnosis of tick-borne infections.
In Part 2, Dr. Bobbi Pritt discusses the molecular methods available for detection and diagnosis of tick-borne infections.
Presenters: Matt Binnicker, PhD
Presenters: Bobbi Pritt, MD
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Our speaker for this program is Dr. Matthew Binnicker, Assistant Professor of Laboratory Medicine and Pathology, Director of the Infectious Diseases Serology Laboratory and Associate Director of the Mycology/Mycobacteriology Laboratory in the Division of Clinical Microbiology at Mayo Clinic. In Part 1, Dr. Binnicker will provide an overview and case studies of the conventional methods for detection and diagnosis of tick-borne infections. In Part 2, which is also posted, Dr. Bobbi Pritt discusses the molecular methods available for detection and diagnosis of tick-borne infections.
In this presentation, we’ll discuss the conventional laboratory methods for the diagnosis of tick-borne infections, with an emphasis on how serology should be used and interpreted in cases of suspected tick-borne disease.
There are several conventional laboratory methods that are still commonly used in clinical laboratories to assist in the diagnosis of tick-borne infections. These include direct smear microscopy, culture, and serology. I’ll introduce these methods, and review their advantages and limitations by discussing three brief case presentations.
The first case is of a 75 year-old male from Wabasha, MN who presents to his local emergency department with a fever and shortness of breath. The patient works as a deer processor and 2 weeks prior to presentation had returned from a hunting trip in Wisconsin. Laboratory results reveal that the patient is thrombocytopenic and leukopenic, and he subsequently develops progressive respiratory distress. Because of his deteriorating status, the patient is intubated and transferred to the intensive care unit.
As a component of the patient’s evaluation, a blood sample is obtained and sent to the laboratory for direct examination, and it reveals the following. The technologist at the bench identifies a number of cells with a characteristic staining pattern similar to the one shown in the image on this slide. As you might have already guessed, a finding such as this on direct microscopic exam, along with the appropriate exposure history and clinical presentation would be highly suspicious for a diagnosis of human ehrlichiosis or anaplasmosis.
So let’s review the conventional laboratory methods that are used for the diagnosis of ehrlichiosis and anaplasmosis. The first method I’d like to discuss is direct examination of peripheral blood smears. This technique has been used for many years in clinical laboratories, and remains one of the most rapid diagnostic methods in cases of suspected ehrlichiosis. In this test, laboratory technologists examine the peripheral smear for the presence of monocytes or granulocytes containing morulae, which is a latin term for “mulberry.”
As you can see in the images on the right-hand side of this slide, the organisms can be seen in characteristic clusters resembling mulberries. While the identification of morulae in peripheral blood smears is relatively specific, it is very insensitive for human monocytic ehrlichiosis, or HME, which is caused by Ehrlichia chaffeensis. We do see a slightly increased sensitivity in cases of human granulocytic anaplasmosis, or HGA, which is caused by Anaplasma phagocytophilum. However, it is still a relatively rare observation, with only 25 to 75% of smears being positive. I should also mention that empiric antimicrobial treatment may further decrease the sensitivity of smear, so ideally, a blood sample should be obtained prior to the initiation of therapy.
The second conventional method for the diagnosis of ehrlichia and anaplasma is in vitro cultivation, which involves recovery of the organisms from blood or CSF in cell culture. This approach is uncommon, and is limited to a small number of specialty laboratories or state and national health department labs. While in vitro cultivation has high specificity, its use is limited by the fact that it requires the application of unique antibiotic-free cell culture media and highly trained personnel to perform testing, and it may require 2-6 weeks for the organisms to be detected by this method.
This brings us to serology, which is the most commonly used conventional method for the diagnosis of ehrlichia and anaplasma infections. This technique shows high sensitivity for the diagnosis of HME or HGA by demonstrating seroconversion or a 4-fold increase in titers between acute and convalescent phase sera. Serology for these infections is most commonly performed by indirect immunofluorescence antibody, or IFA. IFA is often considered the gold-standard serologic method, but it is labor intensive and requires highly trained personnel to perform testing and interpret the results. In addition, it is subjective and therefore prone to inter-reader and inter-laboratory variability.
The next slide shows a typical positive result by IFA, showing characteristic patterns of apple-green fluorescence throughout the visible field. The highest dilution of the patient sample showing green fluorescence is determined to be the end-point titer and this value is what is most commonly reported.
Overall, the performance of Ehrlichia and Anaplasma serology by IFA is good, with sensitivities typically ranging between 80 and 90% for IgG-class antibodies, while lower sensitivities are observed for IgM. In addition, the specificity of the IgG test is reported to range between 80 and 100%, while there is lower specificity for IgM due to cross-reactivity with other infectious and non-infectious etiologies.
One of the most important factors to consider when interpreting the results of serology in cases of suspected HME or HGA is the time-interval between the onset of clinical illness and when the serum sample is collected for testing. This relationship is shown in the table on this slide, in which we can see that the sensitivity of serology for HME and HGA is very poor, between 20 and 50%, when the specimen is collected during the first week after symptom onset. However, the sensitivity of serology increases significantly during week 2 post onset, and by the third week, serology is positive in greater than 90% of cases of HME and HGA.
Other important factors to consider when ordering and interpreting serology results for Ehrlichia and Anaplasma include the fact that IgG may persist for months to years in exposed patients, and that there may be a high seroprevalence in endemic areas. Because of this, IgG-class antibodies to HME or HGA may be detected in individuals with no clinical evidence of acute infection. Additionally, an analysis of a single, acute-phase serum sample has low sensitivity, and the sensitivity increases significantly when testing is performed on both acute and convalescent phase sera, preferably collected at least 2 weeks apart.
Finally, while the specificity of Ehrlichia and Anaplasma serology is good, particularly for the IgG assays, there is the potential that cross-reactivity may occur between HME and HGA serologic tests. Because of this, testing for both may assist in the interpretation, as the etiologic agent usually shows the higher endpoint titer, for example, in cases of Anaplasmosis, we typically see that the HGA titer is greater than the HME titer, often times by at least several fold. When IgM testing is performed for HME and HGA, a positive result without an accompanying IgG titer should be interpreted with caution, as false positives can occur due to Rocky Mountain spotted fever, typhus, Q fever, Lyme disease, or EBV, or elevated rheumatoid factor.
Now that we’ve reviewed how methods, such as serology, can assist in the diagnosis of ehrlichiosis and anaplasmosis, let’s turn our attention to case #2. This is a case of a 4-week old infant who was seen at a Children’s hospital in Rhode Island. The infant received blood transfusions from at least 7 donor units, and within days, the child developed fever, anemia and thrombocytopenia. A peripheral blood specimen was obtained from the patient and was submitted to the laboratory for direct examination.
A review of Giemsa-stained thin preparations revealed the following.
As you can see in these images, red blood cells containing multiple, often time pleomorphic ring forms are observed. These findings, as well as the clinical presentation of the patient would be consistent with a diagnosis of transfusion-transmitted Babesiosis.
As was performed in this case, the examination of Giemsa-stained thin blood smears is the most direct and inexpensive diagnostic approach in cases of Babesiosis. However, multiple smears may be required if there is low parasitemia. In addition, the trophozoite forms of Babesia may be confused with the ring trophozoites of Plasmodium as you can see in the images at the bottom of the slide. So it is important that slides be reviewed by trained personnel, and that the travel and exposure history be examined when interpreting the results.
There are several distinguishing features that can help in differentiating Babesia and Plasmodium when examining thin blood smears. These include the fact that Babesia trophozoites are variable in size, ranging from 1 to 5 microns. Second, multiply infected erythrocytes are more common in Babesiosis. And finally, when tetrads of merozoites, also known as the maltese cross form, are seen, this is pathognomonic for Babesia. However, this is infrequently observed.
Serology may be used to assist in the diagnosis of Babesiosis. The most common serologic approach for Babesia is immunofluorescence, with positive results showing patterns of apple green fluorescence as seen here.
I think the overall utility of serology in cases of Babesiosis is limited, but it may be helpful in cases of low parasitemia that may be missed by blood smear examination. As we discussed with Ehrlichia and Anaplasma, the sensitivity of Babesia serology is low in the first week following the onset of symptoms, but the sensitivity increases during the later stages of disease.
Several factors to consider when interpreting the results of Babesia serology are that an IgG titer of greater than or equal to 1:64 is generally considered positive, but a low titer may be seen in patients from endemic areas without disease, and the results may be interpreted as simply past, asymptomatic exposure.
In contrast, IgG titers of 256 or greater are generally considered to correlate more closely with acute infection, but a comparison of acute and convalescent phase sera is recommended when diagnosing acute disease using serology. It’s important to note that based on current data, the correlation between titer and disease severity is poor, and it is not generally recommended to use titers to monitor a patient’s response to therapy. This is due to the fact that while titers may drop to negative within 6-12 months of infection, titers can also persist for years despite adequate therapy and resolution of symptoms.
The third and final case that I’d like to review in this presentation is of a 15 year old boy who was in his normal state of good health when he developed fever, fatigue and myalgias of 3 days duration. His history was notable in that he had recently returned with his family from a vacation in northern Minnesota. The boy’s mother indicated he had exposure to ticks and mosquitos during the trip. Due to the patient’s clinical presentation and recent travel history, the care provider suspects possible Lyme disease.
When evaluating a patient with suspected Lyme disease, the diagnosis is often clinical, and is based on symptoms and objective clinical findings. The most characteristic manifestation is that of erythema migrans, also known as a target lesion or bull’s eye rash. Erythema migrans is believed to occur in 60-75% of patients with Lyme disease. Other clinical findings may include facial palsy or arthritis.
There are several situations where laboratory testing is not recommended. The first situation is when a patient presents with an appropriate exposure history and a “classic” erythema migrans rash. In this situation, it is typically recommended to bypass laboratory testing and treat the patient according to approved guidelines. The second scenario where lab testing should not be performed is when a patient lacks symptoms, an appropriate exposure history, or is from a nonendemic region and has not recently visited an endemic area.
When laboratory testing is performed, there are several standard methods that can be used. The first is culture, with the optimal specimens including tissue obtained by biopsy of the EM rash, or synovial fluid collected from an affected joint. Although a positive culture for Borrelia burgdorferi is diagnostic, recovery of this organism in culturemay take from 4 days to several weeks, and culture is usually performed only at specialized, reference or government health department laboratories.
The most common approach for the laboratory diagnosis of Lyme disease is through the use of serology, in which methods such as enzyme immunoassay or Western blot are often performed.
Now that we’ve reviewed some of the standard laboratory methods that are available for the diagnosis of Lyme disease, let’s return to our patient case for an update. A serum sample was submitted for Lyme serology, and the results of the screening enzyme immunoassay were positive. The physician receives the report of the screen EIA and questions “Is this diagnostic for Lyme disease?” To answer this question, I’d like for us to review the algorithm recommended by the Centers for Disease Control for Lyme serologic testing.
Currently, the CDC recommends a “2-step” approach for the serologic diagnosis of Lyme disease. In the first tier of testing, the specimen is tested by a highly sensitive screening assay, such as an EIA. If the result of the screening EIA is negative, the report should go out as Negative, and no further testing is required with the present specimen. However, in the case of short disease duration, it may be beneficial to submit a follow-up specimen in 7-14 days if clinically indicated. If, on the other hand, the result of the screening assay is positive or equivocal, the specimen should then be tested by Western blot for IgM and IgG-class antibodies.
Currently, the criteria for determining whether a Lyme Western blot is positive or negative depends on the total number of diagnostic bands determined to be present on the test strip. For instance, an IgM Lyme Western blot is positive if at least 2 of the 3 diagnostic bands are present. And for IgG, the laboratory must observe at least 5 of the 10 diagnostic bands for a positive result. Specimens meeting these criteria are reported as positive, and these results would support a clinical suspicion of Lyme disease. If these criteria are not met, the result is reported as negative and no further testing is required, except in the setting of short disease duration. If this is the case, a follow-up specimen can be tested in 1-2 weeks if clinically indicated.
I just want to emphasize one additional point regarding Lyme WB before we move on. The criteria for Lyme WB were established to guide the interpretation of serum samples, and should NOT be applied when interpreting cerebrospinal fluid serology results. To assist in the diagnosis of neuroborreliosis, we recommend that the CSF sample and a companion serum sample be submitted for Lyme antibody index testing, which is designed to assess whether intrathecal antibodies to Borrelia burgdorferi are present in CSF. Although Lyme serology is a reliable method to assist in the diagnosis, there are several important limitations that we should discuss.
Regarding the screen enzyme immunoassay, the sensitivity of the assay may vary with the disease stage. As we can see in this table, the sensitivity of the screening EIA in stage 1 of the disease may be as low as 70-75%. In contrast, when testing is performed during stages 2 and 3 of Lyme disease, the sensitivity increases to greater than 90%. An additional point to consider is that early administration of antibiotics may weaken or delay the antibody response, and the sensitivity of EIA may be decreased. As we’ve already reviewed, specimens testing positive or equivocal by the screening method should be tested by Western blot.
Lyme Western blot assays test for IgM and IgG-class antibodies to Borrelia burgdorferi.
Western blot should be considered a supplementary serologic approach, and not confirmatory, due to the potential for false positives and inter- and intra-laboratory result variability with this method. And as we’ve discussed, the results of Western blot should be interpreted according to the current CDC criteria, and testing by Western blot is not recommended in those patients testing negative by screen EIA.
As with the screening tests, the sensitivity of Lyme Western blot is also dependent on the stage of the disease when testing is performed. During stage 1, Western blot may demonstrate a sensitivity of only 20 to 50%, with the majority of antibody detected being of the IgM-class. In stages 2 and 3 of the disease, the sensitivity of Western blot increases to 70-100%, and we typically observe a switch in prevalence of antibody to that of the IgG-class.
It is important to point out that although a positive IgM Western blot generally indicates recent infection, IgM-class antibodies may persist for months to years following infection in some cases, and because of this, a positive IgM does not always serve as an indicator of recent, acute infection.
If we now return to our patient case, you’ll remember that the screen EIA was positive, so the testing laboratory appropriately reflexed the specimen to Western blot for supplementary testing. The results of the Western blot were as follows: The IgM was interpreted as positive with 2 of the 3 diagnostic bands present, and the IgG was reported as negative, but with 3 of the 10 diagnostic bands present. A common question following a result profile such as this is “Whether any further follow-up testing should be performed to support the diagnosis?”
For those patients testing positive by IgM Western blot alone, the general recommendation is to repeat serologic testing on a new specimen after 14-21 days. After this time, we should be able to demonstrate seroconversion of IgG, which provides strong supportive evidence of infection.
A few more points regarding Lyme Western blot: If testing is performed within 4 weeks of symptom onset, it is recommended that the results of both IgM and IgG Western blot be used in the interpretation. However, in patients with disease duration of greater than 1 month, the results of the IgM Western blot should not be considered diagnostic, and a positive IgM alone should not be used to diagnose infection due to the increased possibility of false positive results during the later stages of disease.
Returning to our patient case, a second serum specimen was submitted 10 days later for follow up Lyme serology. The result of the screening EIA was again positive, and Lyme Western blot was determined to be positive for both IgM and IgG. These laboratory results provided strong supportive evidence that this patient had been recently infected with Borrelia burgdorferi.
The laboratory diagnosis of Lyme disease is most commonly achieved by serologic testing, which should be limited to those persons with an appropriate exposure history and objective clinical findings. When serology is ordered, testing should be performed using the 2-step approach as recommended by the CDC, in which specimens are first tested by a highly sensitive screening method, such as EIA, with all screen-positive or equivocal specimens being tested by Western blot. And finally, the results of Lyme serology should be used to support a clinical diagnosis in those patients being evaluated for Lyme disease.
In conclusion, conventional laboratory methods for the diagnosis of tick-borne infections include culture, direct smear examination and serology. It is important to remember that serology is generally insensitive during the acute-phase of tick-borne infections, so other testing, such as real-time PCR is commonly recommended. And the final point that I’d like to leave you with, is that the clinical findings and exposure history of your patient should drive the laboratory tests that are ordered. In other words, always think pretest probability!