We evaluate the consequences of processing alcohol-fixed tissue in a processor previously used for formalin-fixed tissue. Biospecimens fixed in PAXgene Tissue Fixative were cut into three pieces then processed in a flushed tissue processor previously used for formalin-fixed, paraffin-embedded (FFPE) blocks (neutral buffered formalin [NBF](+ve)), a formalin-free system (NBF-ve), or left unprocessed. Histomorphology and immunohistochemistry were compared using hematoxylin/eosin staining and antibodies for MLH-1, Ki-67, and CK-7. Nucleic acid was extracted using the PAXgene Tissue RNA/DNA kits and an FFPE RNA extraction kit. RNA integrity was assessed using RNA integrity number (RIN), reverse transcription polymerase chain reaction (RT-PCR) (four amplicons), and quantitative RT-PCR (three genes). For DNA, multiplex PCR, quantitative PCR, DNA integrity number, and gel electrophoresis were used. Compared with NBF-ve, RNA from NBF+ve blocks had 88% lower yield and poorer purity; average RIN reduced from 5.0 to 3.8, amplicon length was 408 base pairs shorter, and Cq numbers were 1.9-2.4 higher. Using the FFPE extraction kit rescued yield and purity, but RIN further declined by 1.1 units. Differences between NBF+ve and NBF-ve in respect of DNA, histomorphology, and immunohistochemistry were either non-existent or small in magnitude. Formalin contamination of a tissue processor and its reagents therefore critically reduce RNA yield and integrity. We discuss the available options users can adopt to ameliorate this problem:

Background: A formal method validation for biospecimen processing in the context of accreditation in laboratories and biobanks is lacking. A previously optimized stool processing protocol was validated for fitness-for-purpose for downstream microbiome analysis. Materials and Methods: DNA extraction from human stool was validated with various collection tubes, stabilizing solutions and storage conditions in terms of fitness-for-purpose for downstream microbiome analysis, robustness, and sample stability. Acceptance criteria were based on accurate identification of a reference material, homogeneity of extracted samples, and sample stability in a 2-year period. Results: The automated DNA extraction using the chemagic™ Magnetic Separation Module I (MSM I) extracted 8 out of 8 bacteria in the ZymoBIOMICS® Microbial Community Standard. Seven tested stabilizing solutions (OMNIgene®•GUT, RNAlater®, AquaStool™, RNAssist, PerkinElmer SEB lysis buffer, and DNA Genotek's CP-150) were all compatible with the chemagic MSM I and showed no significant difference in microbiome alpha diversity and no significant difference in the overall microbiome composition compared to the baseline snap-frozen stool sample. None of the stabilizing solutions showed intensive polymerase chain reaction (PCR) inhibition in the SPUD assay. However, when we take into account more stringent criteria which include a higher double-stranded DNA yield, higher DNA purity, and absence of PCR inhibition, we recommend the use of OMNIgene•GUT, RNAlater, or AquaStool as alternatives to rapid freezing of samples. The highest sample homogeneity was achieved with RNAlater- and OMNIgene•GUT -stabilized samples. Sample stability after a 2-year storage in -80°C was seen with OMNIgene•GUT -stabilized samples. Conclusions: We validated a combination of a stool processing method with various collection methods, suitable for downstream microbiome applications. Sample collection, storage conditions and DNA extraction methods can influence the microbiome profile results. Laboratories and biobanks should ensure that these conditions are systematically recorded in the scope of accreditation.

DNA extracted from formalin-fixed, paraffin-embedded tissue sections is often inadequate for sequencing, due to poor yield or degradation. We optimized the proteinase K digest by testing increased volume of enzyme and increased digest length from the manufacturer’s protocol using 54 biospecimens, performing the digest in centrifuge tubes. Doubling the quantity of proteinase K resulted in a median increase in yield of 96%. Applying the optimized proteinase K protocol to sections deparaffinized on microscope slides generated a further increase in yield of 41%, but only at >50,000 epithelial tumor cells/section. DNA yield now correlated with (χ 2 = 0.84) and could be predicted from the epithelial tumor cell number. DNA integrity was assayed using end point multiplex PCR (amplicons of 100–400 bp visualized on a gel), quantitative PCR (qPCR; Illumina FFPE QC Assay), and nanoelectrophoresis (DNA Integrity Numbers [DINs]). Generally, increases in yield were accompanied by increases in integrity, but sometimes qPCR and DIN results were conflicting. Amplicons of 400 bp were almost universally obtained. The process of optimization enabled us to reduce the percentage of samples that failed published quality control thresholds for determining amenability to whole genome sequencing from 33% to 7%.

DNA extracted from formalin-fixed, paraffin-embedded (FFPE) tissue blocks is amenable to analytical techniques, including sequencing. DNA extraction protocols are typically long and complex, often involving an overnight proteinase K digest. Automated platforms that shorten and simplify the process are therefore an attractive proposition for users wanting a faster turn-around or to process large numbers of biospecimens. It is, however, unclear whether automated extraction systems return poorer DNA yields or quality than manual extractions performed by experienced technicians. We extracted DNA from 42 FFPE clinical tissue biospecimens using the QiaCube (Qiagen) and ExScale (ExScale Biospecimen Solutions) automated platforms, comparing DNA yields and integrities with those from manual extractions. The QIAamp DNA FFPE Spin Column Kit was used for manual and QiaCube DNA extractions and the ExScale extractions were performed using two of the manufacturer's magnetic bead kits: one extracting DNA only and the other simultaneously extracting DNA and RNA. In all automated extraction methods, DNA yields and integrities (assayed using DNA Integrity Numbers from a 4200 TapeStation and the qPCR-based Illumina FFPE QC Assay) were poorer than in the manual method, with the QiaCube system performing better than the ExScale system. However, ExScale was fastest, offered the highest reproducibility when extracting DNA only, and required the least intervention or technician experience. Thus, the extraction methods have different strengths and weaknesses, would appeal to different users with different requirements, and therefore, we cannot recommend one method over another.

OBJECTIVES:To evaluate the stability of RNA and microRNA (miRNA) in PAXgene-fixed paraffin-embedded tissue blocks after 7 years' storage.METHODS:RNA and miRNA were extracted from PAXgene-fixed paraffin-embedded (PFPE) blocks in 2009 then stored at -80°C. Seven years later, RNA and miRNA were again extracted from the same blocks. RNA and miRNA integrity in the 2009 and 2016 extractions were compared using RNA integrity number (RIN), paraffin-embedded RNA metric (PERM), reverse transcription polymerase chain reaction (RT-PCR) for different amplicon lengths, and quantitative RT-PCR (qRT-PCR) for three mRNA and three miRNA targets.RESULTS:In PFPE blocks, mRNA was poorer in 2016 extractions compared to the 2009 extractions in all blocks and all assays applied, with transcripts degrading at different rates in the same blocks. For miRNA, qRT-PCR showed no statistically significant differences between 2009 and 2016 extractions.CONCLUSIONS:mRNA in PFPE tissue blocks degrades at room temperature storage over 7 years.

Vacuum centrifugal (SpeedVac) concentration is commonly applied to nucleic acids extracted from formalin-fixed paraffin-embedded (FFPE) sections, but with an unknown impact. We concentrated DNA and RNA from FFPE biospecimens using different time/temperature SpeedVac combinations of up to 30 min concentration at 45 °C, then used spectrophotometry, spectrofluorometry, RIN, PERM, DV200, qRT-PCR, DIN and the Illumina FFPE QC Assay to assess the changes in quantity, purity and integrity induced by the concentration process. We found the effects of SpeedVac concentration to be inconsequential, but an aliquot of elution buffer should be concentrated for use as the blank in spectrophotometry assays.

The ability to take targeted multiple cores from a single frozen biospecimen would enable several research projects to be fueled from one biospecimen, a small piece of tissue to be quality-control tested, and for pathologically-discrete areas of a biospecimen (e.g., tumor, stromal, and normal tissue) to be selectively sampled for comparative analyses. CryoXtract Instruments' CXT350 Frozen Sample Aliquotter can potentially achieve this by producing multiple cores from one cryopreserved biospecimen without thawing either the parent biospecimen or its daughter cores. It therefore has the potential to add significant value to a tissue banking workflow. We have evaluated its performance while using 614 cores from fecal, liver, kidney, lung, heart, and colon biospecimens. Coring densities of up to five complete and four fragmentary cores per cm(3) are achievable using 3 mm coring probes. Median core weights for tissue were 14.1-17.2 mg (depending on tissue type) and cores ≤325 mg could be taken from fecal biospecimens (depending on the fill-depth of the tube). The coefficient of variation for multiple cores taken from a fecal biospecimen was 11.7%. Between-sample contamination did not occur. RNA Integrity numbers and qRT-PCR analysis demonstrated that coring induced a statistically significant impact on RNA quality that was inconsequential in magnitude and in our view does not represent a barrier for the effective utilization of the technology.

The optional RNase digest that is part of many DNA extraction protocols is often omitted, either because RNase is not provided in the kit or because users do not want to risk contaminating their laboratory. Consequently, co-eluting RNA can become a "contaminant" of unknown magnitude in a DNA extraction. We extracted DNA from liver, lung, kidney, and heart tissues and established that 28-52% of the "DNA" as assessed by spectrophotometry is actually RNA (depending on tissue type). Including an RNase digest in the extraction protocol reduced 260:280 purity ratios. Co-eluting RNA drives an overestimation of DNA yield when quantification is carried out using OD 260 nm spectrophotometry, or becomes an unquantified contaminant when spectrofluorometry is used for DNA quantification. This situation is potentially incompatible with the best practice guidelines for biobanks issued by organizations such as the International Society for Biological and Environmental Repositories, which state that biospecimens should be accurately characterized in terms of their identity, purity, concentration, and integrity. Consequently, we conclude that an RNase digest must be included in DNA extractions if pure DNA is required. We also discuss the implications of unquantified RNA contamination in DNA samples in the context of laboratory accreditation schemes.

RNA extractions are potentially compromised in terms of both yield and quality by ribonucleases (RNases). The pungent and toxic reducing agent β-mercaptoethanol (β-ME), therefore, is commonly added to the biospecimen’s lysis buffer to aid in RNase deactivation. Using different tissue types (liver tissue, kidney tissue, and cell pellets), extraction kits (RNeasy Mini Kit, Illustra RNA Spin Mini Kit, and PureLink Mini Kit), RNA quality assays (RNA integrity numbers [RINs] and quantitative real-time polymerase chain reaction [qRT–PCR]), yield assessments, and in vitro functional RNase assays (RNaseAlert Kit), we demonstrate that β-ME should be replaced by the less toxic dithiothreitol (DTT) alternative.