Tuesday 29 July 2014

IS RETESTING REALLY WORTHFULL IN CLINICAL LABORATORY?



IS RETESTING REALLY WORTHFULL IN CLINICAL LABORATORY?
Since the early 1970s, laboratory medicine specialists have used computer technology and automation to identify and confirm critical laboratory values.(1,2) The historic practice in clinical laboratories has been to automatically repeat laboratory values that are above or below a critical threshold or that trigger other automated “repeat rules” such as a delta check. These practices were established when laboratory instruments were far less reliable than today,(3) yet they persist in many laboratories (including ours). In fact, recent studies show that analytic issues account for only 8% to 15% of clinical laboratory– related errors, with preanalytic and postanalytic errors representing 85% to 92% of all errors.(4,5) Contemporary laboratory instruments use numerous safeguards in their hardware and software to improve the accuracy and reliability of results.
A recent summary of data from a College of American Pathologists (CAP) Q-Probes survey suggests 61% of laboratories still repeat testing for critical chemistry values.(6) The survey also suggests that laboratory test repeat practices have the potential to delay reporting by 10 to 14 minutes and waste resources without significantly preventing analytic errors.6 Recent studies have shown that the practice of repeating tests with critical laboratory values or other results that trigger automated repeating may not be necessary with today’s clinical laboratory automated analyzers.(3,7) A small study examined a total of 580 repeated tests for potassium, glucose, platelet count, or activated partial thromboplastin time and found that the repeated value was within the acceptable difference for 95.3% of the critical value tests repeated and 97.6% of all repeated tests.(3) Another study examined 5 different hematology/coagulation tests and 500 consecutive critical results and the repeated test values for each of the 5 tests. By using their internal definition of acceptable error, Toll et al7 found that 0% to 2.2% of the repeated values for these tests were outside of their acceptable criteria. They concluded that repeated testing for critical values did not offer an advantage or provide additional benefit in hematology and coagulation settings.(7) Neither of these studies examined the time delays in reporting the critical values that repeated testing invariably causes.
The CAP recently published the results of a survey of 86 laboratories, each of which reviewed 40 critical test results from 4 tests at their institutions.(8) The study found that 61% of laboratories always repeated critical results and that the median delay in reporting as a result of repeated testing was 10 to 14 minutes in most laboratories and 17 to 21 minutes in 10% of the laboratories.(6,8) Based on the findings of these studies and the Q-Probes survey, we examined the results of our repeated testing. For more than 20 years, we have repeated all tests in an automated manner when the initial values exceed the AMR of the test (high or low), the result is a critical value, the result fails a delta check, or the value exceeds a preset “review” limit. All of the instruments used are directly interfaced to our laboratory information system, and results are autoverified when all rules are satisfied. When one of the aforementioned flags occurs, results are not autoverified and are automatically repeated by the instrument. For initial values above or below the AMR of the method, the test is repeated on dilution (high values) or the sample is examined for being a short sample (below AMR) and repeated using a special “short sample” cup if necessary. For initial values within the AMR, the technologist reviews the repeated result, and if the result is in agreement, the initial value is verified manually by the technologist. If the difference between the initial and repeated results exceeds the 2 SD range of the quality control sample closest in concentration to the test sample or is greater than 10% (whichever is greater), the test is performed a third time, and the average of the 2 results that agree is reported.
Based on experience and the reports discussed, it is hypothesized that the vast majority of repeated values from clinical chemistry laboratory would agree with the initial value and that there may be only limited benefit in continuing such frequent repeat analyses. It is also hypothesized that repeated analysis for critical values is delaying notification of caregivers about these critical results. The definition of what constitutes a significant difference between repeated values is variable. Allowable error can be defined by biologic variability,(9,10) subjective opinion of what is clinically significant, clinician survey, or regulatory requirements. CAP/CLIA criteria for allowable error should be chosen because they are understood by all and are the criteria by which proficiency testing is judged in the United States. This decision can be questioned because there is not a clear rationale for some of these criteria, which may lead to some of the differences we observed in the number and frequency of errors we identified for different tests. For example, the CAP/CLIA criterion for sodium is ± 4 mmol/L (~ ± 2.8%), whereas the criterion for calcium is ± 1.0 mg/dL (~ ± 10.0%) It could be argued that the former is clinically insignificant, whereas the latter is clinically significant. It is also possible that the differences in the magnitude of what is considered allowable error led to our finding of considerably more errors for repeated sodium testing within the AMR than for calcium.
It is interesting that when the data for these 2 tests is examined closely, this is not the case. There is only 1 additional calcium repeated result of the 3,079 results within the AMR in which the difference is between 0.5 and 1.0 mg/ dL. In contrast, for the 31 errors within the AMR for sodium, 19 exhibited a difference between the initial and repeated tests that exceeded 10 mmol/L. Results below the AMR (linear low) may be due to “short sampling” or other preanalytic or analytic error. Results above the AMR are repeated on dilution to obtain a final result, and the absolute or percentage differences from the repeated results are most frequently greater than the CAP/ CLIA-derived allowable error. Clearly, when initial results are outside the AMR, repeated testing will continue to be necessary.
The results of several studies for repeated testing when automated chemistry testing; the repeated testing is unnecessary and delays the reporting of results, which is a particularly important problem for critical values. Findings of these studies also suggest that for some tests such as sodium and pO2, repeat testing may be necessary to detect some large errors in the initial result. The reasons for these errors are unclear at this time and may require prospective evaluation. Finally, while it may not be necessary to repeat the analysis for samples that trigger a delta check flag, it will still be necessary for technologists to check the identity and feasibility of the result for the previous sample. Delta checks provide a means for identifying mislabeled samples, sample integrity problems as a result of preanalytic problems, and random analytic errors. While all these studies strongly suggests that random analytic errors are rare, it does not address the first 2 causes of a delta check flag, and these will need to be investigated by the laboratory. The delays observed in reporting critical values that result in repeated testing were similar to those described by survey participants in the Q-Probes study.(6,8) It is not surprising that the delays for blood gases were shorter than the delays for other chemistry tests because the analytic time is much shorter. However, the median delays observed for tests such as potassium and glucose are far greater than the actual analytic time. This is most likely due to a technologist taking time to review results, determine if a third test is necessary, and making a decision about manual verification of the final result. Weaknesses of these studies are that the data are from a single laboratory and only several types of automated analyzers. The error rates may vary depending on instrumentation, quality assurance practices, or other variables of individual laboratories. For example, because using multiple instruments performing the same test (eg, 7 Roche Modular P units), calibrations is not accepted if the quality control samples are outside of 1 SD, which may minimize error rates.

Conclusion
Finally, the number of repeated tests observed for the immunoassay and therapeutic drug monitoring categories may be too small to make firm conclusions. Nevertheless, these results suggest that repeat testing for many automated chemistry tests, including critical values, can be stopped and should also serve as a catalyst for other laboratories to examine the value of their repeat testing practices. Doing so can improve patient care by delivering critical values more rapidly and could potentially save 2% to 3% of reagent costs for many tests.

References
1. Lundberg GD. When to panic over abnormal values. MLO Med Lab Obs. 1972;4:47-54.
2. Lundberg GD. Managing the Patient-Focused Laboratory. Oradell, NJ: Medical Economics Books; 1975.
3. Chima HS, Ramarajan V, Bhansali D. Is it necessary to repeat critical values in the laboratory? Lab Med. 2009;40:453-457.
4. Goswami B, Singh B, Chawla R, et al. Evaluation of errors in a clinical laboratory: a one-year experience. Clin Chem Lab Med. 2010;48:63-66.
5. Carraro P, Plebani M. Errors in a stat laboratory: types and frequencies 10 years later. Clin Chem. 2007;53:1338-1342.
6. Paxton A. Critical value repeats: redundancy, necessity? CAP Today. December 2010;24:1.
7. Toll A, Liu JM, Gulati G, et al. Does routine repeat testing of critical values offer any advantage over single testing? Arch Pathol Lab Med. 2011;135:440-444.
8. Lehman CM, Howanitz PJ, Karcher DS. QP102—Utility of Repeat Testing of Critical Values Data Analysis and Critique. Q-PROBES. Northfield, IL: College of American Pathologists; 2010:1-12.
9. Ricos C, Alvarez V, Cava F, et al. Current databases on biological variation: pros, cons and progress. Scand J Clin Lab Invest. 1999;59:491-500.
10. Fraser CG. General strategies to set quality specifications for reliability performance characteristics. Scand J Clin Lab Invest. 1999;59:487-490.

Monday 28 July 2014

AUTOMATION IN SEROLOGY



AUTOMATION IN SEROLOGY
Serology is the scientific study of serum. In medicine, it refers to the diagnostic identification of antibodies in the serum obtained from a patient’s blood sample. In practice, it has many applications. Microbiology, specifically serology, has stubbornly resisted efforts at automation. The reasons are multiple, from lack of space to remote proximity to the main lab, from complexity of procedure to dedicated versus shared FTEs. Serology will always have some level of manual testing involved. The question is how to minimize manual procedures without compromising the quality of results. The upside is potentially huge; as one of the most labor-intensive areas in the lab, automation offers a way to reduce FTEs that are increasingly in short supply. Concomitantly, it holds the promise of significantly
reducing turnaround time and preventing life-threatening errors through sample consolidation; eliminating sample splitting, automating sample handling, and speeding results notification. There is also the advantage of increasing the serology lab’s capacity; as the volume of infectious disease (ID) testing grows, the need to process more samples with reduced resources is becoming acute. The solution may actually lie within the emerging field of microautomation, where serology platforms are combined with sophisticated automation systems. It goes beyond front-end sample handling; it requires an elevated level of automation intelligence to merge autonomous characteristics of different platforms into a unified whole.Serology has traditionally been served by Elisa-based micro-titer plate (MTP) manual or semi-automated benchtop systems. With the expansion of ID testing portfolios on mainframe and dedicated immunoassay analyzers, the opportunity to consolidate has improved. The problem is that no one platform has all the requisite tests.

Benefits of Automation:

Turnaround Time (TAT)
Automation reduces the turnaround time required to report results through several areas. In the instance of this study, the TAT decreased to less than 1 day for 96% of the workload. Every request arriving prior to 4:00 PM is now completed same day. Previously, accessions coming in after 1:00 PM would not be done until the following day. Turnaround time savings also translates into increased quality metrics and physician satisfaction. At MCA, TAT dropped by over 24 hours for over 30% of the tests. The reduced TAT can accelerate patient care pathways, improving patient care.

Labor
Automation dramatically reduces labor elements such as sample handling, sample splitting, interventions, and results reporting.

Workflow Mapping
The most obvious impact from automation and workstation consolidation is improved workflow in the serology laboratory. Since serology is often not physically located near the main laboratory, hooking platforms onto the automation line is not Practical. Workflow mapping can show significant reductions in the number of human steps required to process the workload. This translates to reduced labor and turnaround time (TAT).

Productivity
Improved productivity is defined as increasing output relative to a fixed input, such as labor or resources. Productivity is measured from the individual technologist’s perspective, the overall lab’s perspective, and the Relative Productivity Index (RPI).

Technologist Productivity
Increasing productivity for the technologist means increasing the number of tests that each technologist produces; therefore, reducing the number of FTEs while holding test volume constant reflects increased technologist productivity. Conversely, increasing the number of tests while holding the number of FTEs constant also results in increased FTE productivity

Laboratory Productivity
Productivity gains are found in increased capacity utilization for testing in the lab. By adding automation, the inherent capacity of each system is able to be maximized both in terms of FTEs and instruments having more capacity. As more tests are able to be processed, a.k.a. produced, overall lab productivity goes up. It is possible for a lab to be more productive without the FTE productivity increasing (i.e. more tests are produced due to increased demand, but the number of techs also increases, so the productivity per tech stays the same). It is also possible for the FTEs to be more productive without the lab productivity increasing (i.e. there are less FTEs but the total volume of testing that the lab produces does not change.)

Quality
There are many elements that define quality, but several key areas include reproducibility, lack of repeats, and minimized human error rates. The benefits of automation include elimination of repetitive tasks, such as reduced sample splitting and pipetting, and a highly reproducible process, with minimal direct interaction.

Sample Handling
In the central area for sample processing, human operators only have to open the sample drawer, load the sample tubes, and close the door. From a LEAN perspective, this has a major impact on non–value added tasks, while significantly reducing the potential for human error. Anyone in the lab can operate the system with no need to dedicate the highest skill level technician for daily routine use. A laboratory supervisor with knowledge of software, adjustment, and troubleshooting is enough to ensure smooth operation of the system.

Sample Splitting
Because the automated system uses a primary tube, there are fewer errors than with a manual approach. No sample splitting is required eliminating the potential of technicians pipetting a sample into the wrong tube. As a result, there are fewer manual errors with a reduced need to repeat tests; blood draws are kept to a minimum. This minimizes the amount of tubes, labels, and pipettes that need to be purchased.

Cost
The impact of the efficiency, productivity and quality improvements translates to significant savings for the serology lab. Savings are realized not only in supplies and labor, but in reduced repeats and sendouts. In addition, the improved capacity utilization increases revenue to offset costs.

Summary
The implementation of micro-automation in the serology lab can bring significant improvements in efficiency, productivity, quality and cost to operations. While traditional automation schemes are not practical in most serology lab settings, the combined effect of high volume immunoassay platforms with large ID portfolios in addition to the unique front-end and sample management capabilities System, enable labs to achieve dramatic improvements in their operations. Automation delivers predictable and consistent service coupled with a reduction in staff

Friday 25 July 2014

REVIEW OF PRANALYTICAL FACTORS TO IMPROVE THE QUALITY OF IHC



REVIEW OF PRANALYTICAL FACTORS TO IMPROVE THE QUALITY OF IHC
Immunohistochemistry or IHC refers to the process of detecting antigens (e.g., proteins) in cells of a tissue section by exploiting the principle of antibodies binding specifically to antigens in biological tissues. IHC takes its name from the roots "immuno," in reference to antibodies used in the procedure, and "histo," meaning tissue (compare to immunocytochemistry). The procedure was conceptualized and first implemented by Dr. Albert Coons in 1941.  Immunohistochemistry (IHC) is a method used to determine the expression of biomarkers in tissue.

Immunohistochemical staining is widely used in the diagnosis of abnormal cells such as those found in cancerous tumors. Specific molecular markers are characteristic of particular cellular events such as proliferation or cell death (apoptosis). IHC is also widely used in basic research to understand the distribution and localization of biomarkers and differentially expressed proteins in different parts of a biological tissue. The histological process, which begins with the acquisition of tissue samples and continues through to the interpretation of IHC or ISH results, can be broadly broken down into three main stages:
v  pre-analytical,
v  analytical and
v  post-analytical.
In this review we are going to discuss about the pre analytical procedures that should be taken into account while preparing section for IHC.

IHC: What can go wrong?
v  PREANALYTIC
Ø Fixation
Ø Selection of markers
v  ANALYTIC
Ø Choice of antibody, retrieval
Ø Use of controls
v  POST-ANALYTIC
Ø Interpretation of immunostaining (+/-)
Ø Interpretation of immmunoprofile

Pre-analytical pitfall: Fixation
Ø  Tissue does NOT fix at 1mm/hr
Ø  mm/hour is permeation time
Ø  Cross-linking of proteins takes much longer
Ø  Optimal fixation: 8 – 24 hours

Fixation
Ø  Small biopsies need as long fixation time as large tissue blocks
Ø  Under-fixation causes more problems than over-fixation


Pre-analytical pitfall:Choice of immunomarker
Ø  Critical
Ø  Should always be based on the morphological differential diagnosis

Fixation modifies the physicochemical state, including redox and membrane potentials, of the tissue, and thereby it changes the reactivity of cellular components with the stain. Consequently, the results of various histological and histochemical staining methods are modified depending on the prefixation used. In addition, many other parameters influence the quality and reliability of immunoreactivities, such as the thickness of histological sections, the dilution range of the antisera used as first layers, and the type or composition of the buffers used for dilution of antisera and of the chromogens (e.g., DAB or FITC), or as the rinsing solution. However, a critical comparison of different fixation media is still missing, although bad fixation quality generally strongly impairs the exact localisation of reaction products in the tissues and organs investigated. An ideal fixation should preserve the original structure of the tissue as good as possible and should be able to provide an equivalent close to the natural state. This demand can be accomplished, for example, by fast penetration of the fixation fluid into the tissue, thus avoiding autolysis and guaranteeing rapid conservation

The pre-analytical stage begins as soon as a piece of tissue is removed from its nutritional source (blood supply) and the time to fixation is critical. Degeneration is caused primarily by autolysis, which is a process of self-digestion by enzymes contained within cells; and this begins immediately. This process is accelerated by increased temperatures. Fixatives are used to stop degeneration, while preserving the structure and integrity of the tissue elements as much as possible.

Fixation:
Tissue preparation is the cornerstone of immunohistochemistry. To ensure the preservation of tissue architecture and cell morphology, prompt and adequate fixation is essential. However, inappropriate or prolonged fixation may significantly diminish the antibody binding capability. The most common fixatives used for immunohistochemistry are the followings:
v  4% paraformaldehyde in 0.1M phosphate buffer
v  2% paraformaldehyde with 0.2% picric acid in 0.1M phosphate buffer
v  PLP fixative: 4% paraformaldehyde, 0.2% periodate and 1.2% lysine in 0.1M phosphate buffer
v  4% paraformaldehyde with 0.05% glutaraldehyde (TEM immunohistochemistry
However, fixation itself introduces artifacts and the ideal fixative would also maintain the structure of all of the epitopes in the tissues. This is not achievable, as the alteration in chemical structure caused by fixation necessarily modifies at least some epitopes. For IHC and ISH procedures it is critical that the tissue does not dry out during any stage of the tissue handling and slide preparation. Drying may cause morphological changes, such as poorly defined chromatin; and subsequently alter the structure of the target particularly along the edge of the tissue. This could inhibit ligand binding and is particularly applicable to small specimens such as endoscopic biopsies. Additionally, dry tissue is more adsorbent, which increases the risk of non-specific or unwanted adsorption of reagents during staining procedures, thereby interfering with interpretation of results.

Influence of the temperature and the duration of fixation
An often forgotten but critical aspect is the temperature during fixation. Whereas fixation in Ca-formol and Bouin`s solution are fairly independent of temperature influences and can be mainly conducted at room temperature. In general, low temperatures retard autolysis, but they also decrease the diffusion rate and thus prolong penetration. In our experience, it can be concluded that it is particularly important to have a temperature of not more than 4°C during the incubation. Even only slightly higher temperatures during this fixation process resulted in a reduced quality of tissue preservation.
Furthermore, the duration of fixation influences fixation quality. Fixing tissue in formalin-based solutions for a time less than 24 hrs generally results in a mixture of formalin and ethanol fixation. The latter aspect happens during postfixative rinsing of the samples in 70% ethanol and during embedding. This means that an interruption of the formalin fixation before it is completed will lead to cross-linking only at the tissue periphery promoting a crust formation. In other words, near the centre coagulation occurs, caused by the ascending ethanol solutions during dehydration, or the centre of the tissue sample remains unfixed including tissue hardening.
Moreover, Ca-formol solutions are very susceptible for over fixation problems. Even a controlled prolonged storing in formaldehyde media may lead to excessive cross-linking and cause irreversible damage of epitopes, which diminishes the immunoreactivity during IHC experiments. This advantage is more relevant regarding Bouin`s fixative that is better suitable for a longer fixation, because it generally produces no over fixation effects. Such quality supports the view of Pol André Bouin, who recommended his solution particularly for embryonic tissues. Other restrictions of fixation quality may occur realizing that many organs are composed of different tissues types, which may include varying structural densities with the consequences of varying penetration times of the fixatives within the organ, producing an only more or less acceptable tissue structure. For example, the esophagus epithelium of mammals contains great amounts of keratins, in contrast to most of the other organs, and it is surrounded by a rather voluminous tunica muscularis. Thus, reduced tissue preservation could be a result of such particular cell or organ stabilizing characteristics. Especially the diminished structural quality of Ca-formol fixed tissue might origin from a slow penetration rate of the fluid in such structures, in comparison to Bouin`s solution. As already emphasised earlier, a mixture of different fixatives is still the best way to achieve relatively high quality tissue preservation by a rather steady progress of solution penetration.


Cold Ischemic Time
Recently, there has been more of a focus on “cold ischemic time” and the impact this may have on IHC and ISH results. The duration of cold ischemia is calculated from when the tissue is removed from the body to when the tissue is placed into fixative. This time should be as short as possible, with published guidelines of one hour or less . The deleterious effects of delayed fixation are increased, decreased or de-localized immunoreactivity. It should be noted that deterioration of an epitope due to ischemia cannot be recovered using antigen retrieval techniques. Relatively little has been published on the ischemic effects for specific antigens or molecular targets which are Class I. Perhaps, a broader understanding of the interrelationship between ischemic time and different targets will be easier, once the recording of ischemic times becomes a part of required documentation for all specimens.

Accessioning and Documentation
When the specimen is received in the laboratory it is ‘accessioned’ and given a unique, traceable number. The documentation (requisition) which accompanies each surgical specimen should include: patient and physician information, date of procurement, clinical information, specimen site and type, collection time, cold ischemic time, type of fixative and duration of fixation. If it is necessary to decalcify a specimen, then that information must also be recorded, including: time in fixative before decalcification, the type of decalcification used, the length of decalcification and any post-decalcification treatment. Part of the sample verification process during accessioning is to confirm that the information on the requisition matches that on the specimen container. The specimen container should have a minimum of two identifiers such as patient name and date of birth.

Grossing
Once a specimen is deemed acceptable, it is examined macroscopically. This is referred to as grossing and it is a critical pre-analytical step which requires proper training. Larger specimens should be ‘bread loafed’ (sliced) into approximately 5 mm sections and placed in 10% NBF. Gauze or paper towel may be placed between the slices to facilitate exposure to the fixative. Care must be taken to handle each type of tissue in a standardized manner and not to physically damage the tissue. Usually, it is necessary to select areas of interest from a larger specimen. These pieces of tissue, or blocks, should be trimmed such that the size does not exceed 20 mm in length and width, or 4-5 mm in depth. Trimmed tissue is then placed into a processing cassette and submerged immediately into the desired fixative (usually 10% NBF). The volume of fixative should be approximately 10 to 20 times that of the specimen . Formalin enters the tissue relatively quickly, but the chemical processes which actually fix the tissue are more time consuming, taking at least 24 hours. When calculating total time in fixative, the time the specimen sits in 10% NBF in the grossing area and on the automated tissue processor must be included.


Tissue Processing
During tissue processing, fixation reagents containing water are replaced by wax (polymer, non-polymer and microcrystalline formulas exist) which is done through a series of passages through increasing concentrations of alcohol, up to 100% (absolute) alcohol. This process is followed by clearing the alcohol from the tissue (for example by using xylene) and replacing it with molten wax. Low melting temperature (45 °C) as opposed to higher melting temperature (65 °C) waxes have been reported to produce better staining results for IHC, particularly in T-lymphocyte staining. Next, the paraffin infiltrated pieces of tissue are embedded to form blocks, which are easily handled, cut and subsequently stored.

Rapid Tissue Processing
There is increasing pressure to shorten turnaround times (TATs) in tissue pathology, so that patients do not have to wait days to receive their pathological diagnoses. However, the laboratory staff still need to ensure that samples are properly fixed (>24 hours in NBF, even for needle biopsies), to make sure that validated IHC and ISH methods are used. As noted above, tests used after alternative fixation and processing must be fully re-validated. This requirement is particularly applicable to the modern rapid tissue processors which employ alternative fixatives and microwave enhanced processing (as well as small specimen size). This combination allows an H&E diagnosis on paraffin sections the same day. Nonetheless, the morphology will differ from routine FFPE processed samples; and IHC and ISH methods will require complete re-validation, as some of these will not need pre-staining antigen retrieval, whether this is of the heating or proteolytic type (personal observation). Section Preparation Generally, unless otherwise specified by a protocol of choice, sections for IHC or ISH are cut at 3 μm, 4 μm or 5 μm. Thicker sections may cause difficulty during staining, and also problems in interpretation due to the multi-layering of cells. After sections are cut they are usually floated on water and picked up onto glass slides that are coated with some adherent material. Sections must lay flat against the glass to prevent lifting during staining or bubble formation, which may trap staining reagents. The more points of adhesion the more likely the tissue will remain fixed to the slide, supporting the need for thinner sections. Some commercially available slides come with a positive charge that attracts the negative charges of tissue proteins. These charged slides are especially effective following formalin fixation of tissues, since formalin blocks amino acids in tissues, rendering the tissue more acidic and therefore more negatively charged. Different manufactures of staining platforms may recommend the use of particular slides to achieve optimal results. As with every other pre-analytical step, cutting and mounting sections onto glass slides, and all steps prior to staining must be standardized. For example, if the slides are to left at room temperature for 15 minutes, in an upright position to allow draining of excess water and then heated in staining rack at 60 °C for 30 minutes prior to staining, this step must be repeated every time sections for IHC or ISH are prepared. Finally, the changes resulting from block and section storage prior to IHC and ISH staining may also affect staining results. For example, it is recommended that sections cut for HER2 testing should not be used if they are more than 6 weeks old.

Dewaxing and Hydration
Wax must be removed completely from the tissue sections, so that aqueous antibodies, molecular probes and detection reagents can penetrate and adhere to the tissue. Traditionally, dewaxing was done by immersing the sections into a dewaxing solution (such as xylene), with or without prior brief heating. This step was followed by absolute and graded hydrating solutions (generally alcohols), until the final solution: water. If xylene is used to dewax sections, approximately 50 slides per 50 mL of xylene is the limit, before it is no longer effective and residual wax begins to cause artifacts in the stained tissue. Today, there are many commercially available staining platforms which include onboard removal of wax and rehydration of the tissue sections. The accumulation of residual wax may be a problem with these instruments, if rinsing is insufficient or if solutions are not replenished regularly.

Conclusion
Patient safety based on accurate interpretation of results depends heavily on this standardization of all pre-analytical variables. Prognostic tests using IHC and ISH are being developed and they will independently forecast clinical outcomes for patients. HER2, ER and PgR are considered predictive markers that influence the selection of patients who will respond more favorably to therapies, emphasizing further the need for standardization. Even if it is not possible to perfectly optimize every pre-analytical step, it is possible to perform each step in the same manner each time it is done. Rigorous adherence to this approach will yield more meaningful results and will, if necessary, facilitate problem solving