Edited by Daniel M. Baer, M.D.
An
interesting array of lab queries gets expert attention
Giardia and Cyclospora PCR
I am trying to detect Giardia lamblia and Cyclospora cayetan- ensis from stool samples extracted by Qiagen stool mini-kit, and used the Weiss and Relman PCR protocols, respectively. I have problems with the product amplification. I tried with positive fecal samples by microscopic and ELISA evaluation and with positive proficiency material without satisfactory results. I do not know if the problem is the protocol, the reagents or the primers. I have the following protocol:
Giardia lamblia: 18S region (182bp) JW1 _ GCGCACCAGGA ATGTCTTGT, JW2 _ TCACCTAC GGATACCTTGT 50 cycles (95C for 1 min, 50C for 1 min, 72C for 1min). Do you have a different protocol for detection for
Giardia lamblia by PCR or any advice?
A:
The traditional way to develop this PCR test would be to first use known
Giardia DNA to establish the optimum conditions for the assay. Then spike negative stool specimens with known
Giardia DNA and repeat the assays. Follow by performing the test on known positive patient stool samples. Concerning your assay, are your stools fixed, and if so, how, and have you extracted all the inhibitors normally present in stool samples? For this, you may need to add a known DNA and perform a PCR test to see if the DNA is amplified in each stool sample indicating that there are no inhibitory substances that would keep the PCR test from working properly. In addition, with the use of as many as 50 cycles, it might be best to perform nested PCR. Try to contact people published in this area to gain more knowledge and troubleshooting options. I would like to thank Dr. Theodore E. Nash, head of the Gastrointestinal Parasites Section, Laboratory of Parasitic Diseases at the National Institute of Health for his insight into this question.
Susan E. Sharp, PhD, DABMM
Director of Microbiology
Kaiser Permanente;
Associate Professor
Pathology Regional Laboratory
Oregon Health Science University
Portland, OR
Calibration verification
Q:
The College of American Pathologists (CAP) requires calibration verification and linearity checks for accreditation. The Clinical Laboratory Improvement Act (CLIA) states calibration verification is only required for modified moderately complex tests. Since CAP does not recognize moderate complexity everything is high complexity I am confused. Are there established tolerances or procedures for in-house calibration verification methods? Is running calibrators as unknowns appropriate for calibration verification?
A:
The regulations are written in government legalese, which to the untrained is a foreign language.1,2 This section (Subpart K, 493.1217) required explanation after it was published. Many people referred to this as a linearity check, while the authors of the regulations insisted it was calibration verification.
Calibration verification is only required on modified moderately complex and high-complexity tests, but if CAP inspects you, it applies to both moderate and high complexity. If a procedure is calibrated with less than three calibrators (most linear single-point methods), then calibration verification must be performed every six months with each new lot of reagents and when major maintenance is done on the instrument. This should consist of at least a zero or minimum standard, a midpoint standard, and a standard with a value at the upper end of the reportable range.
Calibrators can be purchased or made by diluting a specimen with an elevated value and using diluent for the zero standard. What is acceptable verification is not defined in the regulation. You might make a policy of 3 SD of the low control for the minimum standard and 5% of theoretical for others. You must be able to justify your acceptability standards to an inspector. You cannot report patient values (without diluting) that are above or below your lowest or highest calibrator verifier. As to benefit for linearity surveys, a 1995 paper showed that labs with poor calibration evaluations on linearity surveys had higher unacceptable rates on proficiency
testing.3
The relevance and interpretation of calibration, calibration verification, linearity and two newer concepts the analytical measurement range (AMR) and clinically reportable range (CRR) continue to evolve. The highest and lowest calibrators or calibration verifiers (i.e., controls) define the reportable range for patient specimens or the AMR. The AMR can be extended by adding additional calibrators or by diluting the specimens; the result is the
CRR.
Linearity requirements have been eliminated from the CLIA regulations and from the CAP inspection criteria, however, many inspectors continue to feel that linearity studies are a part of good lab practice and should be encouraged. The choice of use of any linearity product lies solely with the lab director. If a lab chooses to continue linearity studies, these studies must fully comply with the calibration/calibration verification requirements of CLIA and/or CAP.
Ronald Laessig, PhD
Sharon Ehrmeyer, PhD
Department of Pathology and Laboratory Medicine
University of Wisconsin
Madison, WI
Ronald D. Feld, PhD
Associate Professor and Director,
Clinical Chemistry Laboratories
University of Iowa Hospitals and Clinics
Iowa City, IA
References
- Passey RB. How to read the Federal Register and other
CLIA-related documents. MLO.1992;24;10:47-51. - Passey RB. Is CLIA understood by clinical
laboratorians? MLO.1994;26;7:54-60. - Lum G, Tholen DW, Floering DA. The usefulness of calibration verification and linearity surveys in predicting acceptable performance in graded proficiency tests. Arch Pathol Lab Med.1995;119:401-408.
D-dimer, FDP and D-dimer quantitation
Q:
We offer D-dimer and FDP
manual tests in the DIC panel in our laboratory. Is it beneficial to offer both tests? Could we replace the manual tests with an automated D-dimer quantitation? Are there any other conditions where the FDP might provide useful information?
A:
An article discussing the
D-dimer test in the diagnosis of venous thrombosis was recently published, and the reader is encouraged to study this review in order to maximize the usefulness of this test in the local
laboratory.1 The D-dimer assay can be used to rule out deep venous thrombosis (DVT) or pulmonary embolism (PE) if levels are not elevated. But many diseases, in addition to DVT or PE and clinical procedures can result in elevated D-dimer levels.
The problems with D-dimer tests have been known for some time, and they persist until the present. A recent article illustrates the complexity of the variety of monoclonal antibody methods (e.g., latex agglutination methods, ELISA and immunoassay methods), which are used. But there is a warning that there may be even more problems in the not-too-distant future, as D-dimer is called to perform in other venues. Again, the importance of local standardization of the testing, including the determination of predictive values, sensitivity and specificity, is stressed. Working closely with the clinicians also can significantly improve the usefulness of D-dimer
testing.2 A helpful list of current references is included.
FDP testing appears to be decreasing in its application to these clinical problems.
John A.
Koepke, MD
Professor Emeritus of Pathology
Duke University Medical Center
Durham, NC
References
- Marlar RA. D-dimer: Establishing a laboratory assay for ruling out venous thrombosis. MLO. Nov, 2002:28-32.
- Titus K. Identity crisis persists – Which D-dimer? CAP Today. 1 January 2003;Vol.17:1,12-16.
Personal protective equipment
Q:
Lab workers have been wearing personal protective equipment or PPE (e.g., gowns, gloves, face shields) for years. Have these safety measures reduced laboratory-acquired infections? Is it necessary to wear gloves in the microbiology lab?
A:
Many lab safety measures are
implemented to protect workers from pathogens transmitted by aerosols (e.g.,
Mycobacterium tuberculosis) or blood and other potentially infectious material (e.g., hepatitis viruses, HIV) because these common microbial agents cause serious infections. Lab workers need to recognize that other less common agents (e.g., bioterrorism agents, such as
Brucella and Francisella tularensis) may cause lab infections. Transmission of the bloodborne viruses usually occurs through needlesticks, cuts or contact of mucous membranes with infected blood or other body fluids.
Since the introduction of hepatitis B virus (HBV) vaccine in 1982, the rate of occupationally acquired HBV disease has fallen from approximately 12,000 cases to 800 cases/year in
1996.1 As of June 2001, only 57 cases of documented occupationally acquired HIV have occurred in the United States (16 were clinical laboratory personnel). Because only a small number of infections occur in any one laboratory and these data are not often reported, it is difficult to document that the use of PPE has reduced the number of laboratory-acquired infections in a specific lab.
The training and education that has occurred through use of these preventive measures including safety-engineered devices and work practices can be measured by other outcome audits (e.g., number of needlesticks accidents), and these data do demonstrate a positive
change.2 One key to reducing lab infections is recognizing the reservoir, routes of transmission and point of entry of an infectious agent, and employing the necessary barriers and techniques to prevent infection (including standard precautions). The concept of standard precautions refers to the idea that all patients and all laboratory specimens should be considered potentially infectious, regardless of the patients diagnosis or presumed infectious
status.1 When handling these specimens, the appropriate PPE should be worn. Gloves should always be worn at the specimen receiving and setup areas, in the TB and virology labs, and whenever contact with potentially infectious or contaminated material may
occur.1 The routine use of gloves when handling bacterial culture plates remains controversial.
David Sewell, PhD, ABMM
Director of Microbiology
Veterans Affairs Medical Center
Portland, OR
References
- NCCLS. Protection of laboratory workers from occupationally acquired infections. Wayne, PA: NCCLS; 2002. M29-A, 2nd ed.
- 2. Fleming DO. Prudent biosafety practices. In: DO Fleming and DL Hunt, eds. Biological Safety: Principles and Practices, 3rd ed., Washington, DC. American Society for Microbiology; 2000: 369-381.
Daniel M. Baer is professor emeritus of laboratory medicine at Oregon Health and Science University in Portland, OR, and a member of MLOs editorial advisory board.
Tips from the Clinical Experts department provides
practical, up-to-date solutions to readers’ technical and clinical
issues from a panel of experts in various fields. Readers may send
questions to Dan Baer by fax, (503) 636-7932; or email, [email protected].
December
2003: Vol. 35, No. 12
© 2003 Nelson Publishing, Inc. All rights reserved.
