We thank our collaborators, Stephen Jones, Nia Charrington, Dr. Dianna Maar, and Dr. Samantha Cooper from the Digital Biology Center (Bio-Rad Inc. CA) for their assay design support; Nezar Rghei and Dr. Moemen Abdalla (Norgen Biotek, Canada) for critical advice when optimizing the RNA extraction protocol; and Shannon Campbell, Scott Thurston, Jeff Fensterer, Shannon Martello and Joellyn Enos for assistance with test requirements and commercial monitoring.

The manufacturers&#39; instructions are followed for the reagents listed below, unless otherwise described. The PCR assays are commercially available products designed to detect ROS1 and RET fusions.
1. Working with RNA in Preparation for Reverse Transcription (RT)-dPCR: Laboratory Best Practices
<ol>
	<li>Create an RNase-free environment when working with RNA.
	<ol>
		<li>Use commercially available sprays designed to inactivate contaminating RNases.</li>
		<li>Use certified RNase-free reagents, tips, and tubes. Use barrier tips for pipettors to prevent introduction of RNases or cross-contamination of samples.</li>
	</ol>
	</li>
	<li>Always wear a laboratory coat to prevent particulates from falling from clothing into your sample. Designate a lab coat specific to use with RNA processing.</li>
	<li>Wear gloves to prevent sample contamination from RNases in skin. Change gloves frequently. 
	Note: Assume laboratory surfaces are contaminated with RNase since they are exposed to the environment. Gloves that contact skin, hair, doorknobs, freezer handles, pens/markers, etc. are assumed to no longer be RNase-free.</li>
	<li>Decontaminate pipettors, benchtops, centrifuges, and other work surfaces with a RNase inactivation spray prior to use.</li>
	<li>If possible, maintain a set of equipment for use with RNA only.</li>
	<li>Minimize disruption of air flow in lab areas when working with RNA samples to prevent particulates from falling into samples or contaminating the work area.</li>
	<li>Store purified RNA at -80 &#730;C.</li>
	<li>Avoid multiple freeze-thaws of RNA samples, as this can cause degradation.</li>
</ol>
2. Generation of Analytic RNA Material for the Positive Controls
<ol>
	<li>Design synthetic DNA using published mRNA sequences for fusion variants of interest10.
	<ol>
		<li>For a given fusion variant, select an mRNA fusion sequence that includes the fusion site plus sufficient length flanking on each side to cover the PCR amplicon.</li>
		<li>Select nucleotide sequences between 50 - 250 nt to mimic the size of circulating RNA captured using platelet-enriched plasma.</li>
		<li>Add a T7 promoter sequence (5&#39;-CAGAGATGCATAATACGACTCACTATAGGGAGA-3&#39;) to the 5&#39; end of the target sequence.</li>
	</ol>
	</li>
	<li>Order synthetic sequences as double stranded deoxyribonucleic acid (DNA) fragments.</li>
	<li>Reconstitute synthetic DNA in Tris-EDTA (TE) buffer to a final concentration of 10 ng/&#181;L.</li>
	<li>Convert 60 ng synthetic DNA to RNA using in vitro transcription.</li>
	<li>Purify RNA transcripts using phenol/guanidine-based reagent12.
	<ol>
		<li>Include DNase I, RNase-free to remove residual template DNA.</li>
	</ol>
	</li>
	<li>Measure concentration of purified in vitro RNA using a commercially available fluorometer with RNA-specific dyes and standards. Ensure the RNA is within the range acceptable for the chosen standards. Dilution may be required.</li>
	<li>Confirm successful transcription by gel electrophoresis using a 2% agarose gel mixed with RNA gel stain and a high range RNA ladder including 50 - 250 nt size range.
	<ol>
		<li>Load 500 ng of each in vitro RNA onto a gel.</li>
		<li>Run the gel at 5 V/cm.</li>
		<li>Visualize single bands using illumination, and document the results.</li>
		<li>Confirm expected transcript size for each of the fusion variants (based on design in step 2.1.2).</li>
	</ol>
	</li>
	<li>Confirm detection of each in vitro RNA by RT-dPCR using matched variant-specific PCR assay (see steps 5 - 8 of this protocol).</li>
	<li>Optional: Prepare an equimolar mixture of in vitro RNA that contains each of the fusion variants and the control gene GUSB.</li>
	<li>If Step 2.9 is performed: Confirm detection of each of the fusion variants included in the control mixture by dPCR using variant-specific PCR assays (see steps 5 - 8 of this protocol).</li>
	<li>Determine desired input concentration for analytic positive controls by testing concentrations ranging from 0.25 to 2.5 fg10. Choose concentration based on desired copy number output.</li>
	<li>Following confirmation, prepare 10 &#181;L single-use aliquots of analytic RNA for use in the positive control (step 4.4) and store at -80 &#730;C.</li>
</ol>
3. Donor Specimens
<ol>
	<li>Collect 10 mL human whole blood specimens in 10 mL blood collection tubes (BCT) containing a cell-free RNA preservative. 
	Note: All human donors shall consent to research use and no donor-specific identifying information shall be collected or used during testing.</li>
	<li>Process whole blood samples within the timeframe specified by the BCT manufacturer.</li>
	<li>Pooled normal human plasma can be purchased from a commercial source for use within the analytic positive control. Prepare single-use, 1 mL aliquots of the pooled normal human plasma and store at -80 &#730;C for use with the positive control (step 4.4).</li>
</ol>
4. Recovery of Circulating RNA from Plasma
Note: It is important to work quickly during this procedure.
<ol>
	<li>Centrifuge whole blood tubes at 200 x g for 20 min.</li>
	<li>Collect up to 4 mL of plasma from centrifuged blood collection tube using a serological pipette. Be careful not to disturb or aspirate the buffy coat layer.</li>
	<li>Isolate circulating RNA using a commercially available kit that can capture exosomes, platelets and cell-free RNA from plasma. Isolate RNA from positive control sample alongside each batch.</li>
	<li>Prepare the Positive Control for each batch of clinical samples as follows:
	<ol>
		<li>Thaw 1 mL pooled normal human plasma aliquot (Step 3.3).</li>
		<li>Thaw 10 &#181;L analytic RNA aliquot (Step 2.12).</li>
		<li>Prepare positive control by adding 10 &#181;L analytic RNA into the normal human plasma sample once ethanol has been added to the plasma lysate.</li>
	</ol>
	</li>
	<li>Elute samples with 100 &#181;L nuclease-free water. Proceed immediately with RNA clean up and concentration.
	<ol>
		<li>Samples may be stored on wet ice, and covered, for up to one hour.</li>
	</ol>
	</li>
	<li>Concentrate RNA using column based method and elute in 9 &#181;L of RNase-free water.
	<ol>
		<li>Proceed immediately to Step 5, or keep samples on wet ice for up to one hour.</li>
	</ol>
	</li>
</ol>
5. Reverse Transcription of RNA to cDNA
<ol>
	<li>Convert concentrated circulating RNA sample to cDNA using a commercially available reverse transcription reaction kit, including random primers (see Table 1 for components). 
	Note: Gene specific primers are optional and can be designed for test variants. Primers are designed based on the target RNA sequence. Use fusion variant sequences from Step 2.1.
	<ol>
		<li>Include no reverse transcriptase control sample and no RNA control sample (see Table 1).</li>
	</ol>
	</li>
	<li>Isolate cDNA from reverse transcription reaction using a commercially available DNA concentrator spin column. 
	Note: This step facilitates removal of enzymes, primers, and free deoxynucleotide triphosphates (dNTPs).</li>
	<li>Use cDNA immediately in PCR reactions or store at -80 &#730;C.</li>
</ol>
6. Digital PCR
Note: This PCR is specific to droplet digital PCR (See Table of Materials).
<ol>
	<li>PCR mix precautions.
	<ol>
		<li>Wear a disposable lab coat and nitrile gloves.</li>
		<li>Use PCR mix reagents in a dedicated reagent preparation area. Do not handle cDNA in the reagent-only preparation area.</li>
		<li>Cover probes while working to protect them from light. Excessive light can photo bleach the fluorescent dye attached to the probe.</li>
		<li>Transport mixes, covered and protected from light, into a separate pre-amplification area before cDNA is to be added.</li>
		<li>Add cDNA to be tested to PCR mix in a PCR clean hood located in pre-amplification area.</li>
	</ol>
	</li>
	<li>Prepare PCR mixes for a final reaction volume of 20 &#181;L according to Table 2.</li>
	<li>Distribute PCR mixes + cDNA to PCR plates. 
	Note: Use of a plate layout as a guide is recommended.</li>
	<li>Cover the plate using a removable plate sealer.</li>
	<li>Centrifuge plates briefly to collect samples at the bottom of the wells.</li>
	<li>Mix on plate shaker on a low setting for 10 s.</li>
	<li>Centrifuge plates briefly to collect sample at the bottom of the well.</li>
	<li>Remove plate sealer. Perform droplet generation for PCR-cDNA mix with either a manual or automated droplet generation system.
	<ol>
		<li>For manual droplet generation, transfer 20 &#181;L PCR mix to sample wells on the droplet generation cartridge. Add 70 &#181;L of droplet generation oil. Cover with rubber gasket and transfer cartridge to manual droplet generator to initiate droplet generation. Following droplet generation, transfer droplets to a fresh PCR plate using tips recommended by the manufacturer. Aspirate and dispense the droplets slowly, over 5 to 6 s each, without touching the opening of the tip to the droplet cartridge or plate.</li>
		<li>For automated droplet generation, seal the plate with a foil seal and transfer to the droplet generator. Ensure all tips, cartridges, and plates are in place before starting droplet generation.</li>
	</ol>
	</li>
	<li>Following droplet generation and transfer of droplets to a fresh PCR plate, seal with a foil plate sealer, and thermal cycle plates using the settings in Table 3.</li>
	<li>After the thermal cycler run is complete, read the plate using a droplet reader. Create a plate layout for reader software that identifies the location of controls, samples, etc., and load into software to start read.</li>
</ol>
7. Data Analysis and Review, and Generation of Results
<ol>
	<li>Analyze plate read results using commercially available software.</li>
	<li>Navigate to the Analyze menu to view two-dimensional (2D) amplitude plots.</li>
	<li>Evaluate the overall quality of the data by examining the droplet data.
	<ol>
		<li>Evaluate data for total accepted event numbers using the Events menu. If there are fewer than 10,000 events per well, carefully evaluate the data for additional problems.</li>
		<li>Check data for aberrant fluorescence amplitudes. Significant amplitude differences and concentration differences between replicate samples indicate poor handling or mixing of samples.</li>
		<li>Make notes of droplet clusters with spray patterns on a 45-degree axis, which is indicative of poor quality droplets or problematic samples.</li>
		<li>Examine Positive Control, No Reverse Transcriptase (No RT), and No RNA Control (NRC) data first. Select all control samples and examine cluster quality by 2D plot. For proper thresholding, a clear separation between droplet clusters should be apparent.</li>
	</ol>
	</li>
	<li>For each assay variant, set the threshold based on control wells.
	<ol>
		<li>Set thresholds on 2D plots using the crosshairs tool to separate the double negative droplet population from the control gene population (labeled with 5&#39;-hexachloro- fluorescein-CE phosphoramidite probe), y-axis, and variant gene population, if present (labeled with fluorescein amidite OR 6-carboxyfluorescein probe), x-axis.</li>
		<li>Sum copies from each replicate well for a single sample.</li>
		<li>Express test results as the number of variant copies detected. 
		Note: To determine the analytic cutoff value for calling a positive or negative sample, run a normal healthy donor sample set (at least 10 individual samples) through the finalized process and establish the cutoff above any detectable background signal for the mutation of interest. Additionally, establish the number of control gene copies required to call a positive or negative result. This control gene cut-off functions as an internal quality control (QC) to evaluate the quantity and quality of each RNA sample that is processed.</li>
	</ol>
	</li>
</ol>
8. Verification of RT-dPCR Reaction Conditions Using Cell-lines (Optional)
<ol>
	<li>To verify detection of fusion variants, use commercially available cell-lines expressing the ROS1 or RET fusion mRNA of interest. Proceed as follows:
	<ol>
		<li>Homogenize flash-frozen cells in a guanidinium-based lysis solution directly from the frozen state. Even brief thawing prior to homogenization can cause RNA degradation and loss.</li>
		<li>Isolate RNA using silica-membrane spin columns designed for RNA.</li>
		<li>Measure the concentration of RNA samples using a fluorometer with RNA-specific reagents and standards.</li>
		<li>Dilute isolated RNA into a background of wild-type RNA from plasma or another commercial source.</li>
	</ol>
	</li>
	<li>Carry out steps from Reverse Transcription of RNA to cDNA, Digital PCR, and Data Analysis and Review, and Generation of Results listed in this protocol to confirm detection of the desired variant.</li>
</ol>

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H.M., L.J., K.A., and G.A.P. are employees of and hold shares in Biodesix, Inc. H.M., L.J., and G.A.P. are co-inventors on a patent application filed by Biodesix, covering a diagnostic test system for the detection of circulating genetic variants in non-small cell lung cancer.

RET and ROS1 rearrangements together make up ~3% of the driver mutations within the NSCLC population18. Although rare, the detection of these genetic alterations is vital. NSCLC patients with these alterations may benefit from targeted therapeutics that specifically inhibit the aberrant kinase activity that results from the onco-protein13. Some such therapies are already approved by the FDA for use in ROS1 positive NSCLC, while others have been demonstrated to be efficacious against RET in clinical trials19.
Digital PCR technology provides sensitivity that is ideal for liquid biopsy applications20. There has been significant adoption of this technology for use with circulating cell-free DNA for the measurement of tumor mutations in patients with NSCLC4,6,21,22,23. In addition to cfDNA, we developed a protocol designed for robust measurement of the most prevalent fusion variants in patients with NSCLC from circulating tumor RNA (Figure 1A)10.
Our established protocol allows for analytic limits of detection down to 0.2% (Figure 2). While the RT-dPCR is exceptionally specific and sensitive, the assays are limited to the panel of known fusion variants that are chosen and multiplexed for detection in the PCR assay. Thus, fusions to be included in multiplexed assays must be carefully selected to ensure proper coverage within the population of patients with NSCLC. We have successfully designed assays for RET and ROS1 that simultaneously detect eight fusion variants resultant from rearrangements of the RET or ROS1 loci and cover 99% and 88% of the RET and ROS1 positive population, respectively (Figure 1B-C)17.
The final test workflow as described in this study includes batch controls to ensure consistency of results. These controls include both a positive analytic standard as well as two negative controls, which together ensure there is no contamination or PCR inhibition occurring within the batch (Figure 3). To ensure robustness of the assay, a study was performed using the batch controls over a 21-day period (Figure 3D,H). These data demonstrate the consistency of the RNA process as established within this protocol.
Good laboratory practices and appropriate RNA handling are key components of ensuring robust and accurate results. Laboratory space and equipment dedicated to use with RNA, cleaning of equipment after each use, using RNase-free reagents and consumables, and applying an RNase inactivation spray to the work space all help to reduce contaminating RNases. Conscientious handling of RNA samples by technicians, including a dedicated lab coat, frequent glove changes, working quickly through the RNA extraction procedure, and keeping samples on ice are of utmost importance to preserving sample integrity. Once RNA has been reverse transcribed to cDNA, the sample is in a more stable form that is less prone to degradation. In addition to practices that support RNA integrity, PCR components and samples should be maintained in segregated areas to prevent cross-contamination that could lead to false positive results. The stock PCR reagents and preparation of PCR master mixes should be kept separate from PCR templates and great care should be taken to segregate amplified template (post-PCR) from all pre-amplified materials including reagents, RNA, and cDNA samples. Finally, proper generation and handling of emulsified PCR mixes prior to amplification is central to maintaining droplet integrity and optimal dPCR conditions. Precautions like these are critical during execution of this protocol to obtain consistent and accurate results. All data should be examined by trained personnel before the release of results to be sure that all QC metrics have been met. In the case of suboptimal results (Figure 4), the batch must be reviewed by technical staff and the Laboratory Director and may require re-processing.
RT-dPCR results can be produced as early as 24 hours from sample receipt and 95% of sample results within the test set used in this study (n = 984) were reported to the ordering physician in less than 72 hours from the time of receipt (Figure 5). This turn-around time provides physicians with much needed molecular information in a time frame that allows initiation of the appropriate therapy. These results are typically available earlier than those obtained using a conventional tissue biopsy. Additional biomarkers for NSCLC and other cancers could be developed using similar circulating RNA-based approaches, and would benefit from the same rapid time-to-results. For example, measurement of the Programmed Death Ligand 1 (PD-L1) mRNA transcript using RT-dPCR could inform physicians about immunotherapy options. There is also a growing interest in the utility of liquid biopsy and dPCR in monitoring for therapeutic efficacy. Earlier indications of resurgence of the tumor using genomic testing for specific variants could allow physicians to adjust treatment regimens before patients are symptomatic by standard of care measures such as imaging24. Protocols such as the one reported in this study are ideal for monitoring due to their non-invasiveness, sensitivity, rapid turn-around time, and cost effectiveness. The assay described herein provides results within 72 hours from sample receipt, with minimal false-positive detection rates, which facilitates rapid treatment decisions and circumvents some limitations experienced with tissue-based testing4.
Our protocol and data demonstrate a robust test system for identifying low abundance RNA variants as well as the potential for blood-based mutation testing in clinical practice. For those patients who do not have an actionable driver mutation identified by rapid targeted liquid biopsy approaches like this one, the addition of more extensive genome and proteome testing from both tissue and blood may provide even broader clinical information to support treatment planning.

Up to 25% of non-small cell lung cancer (NSCLC) patients may not have sufficient tissue available for testing at the time of diagnosis. Even in cases where tissue is available, it may not be of sufficient quantity or quality to perform recommended molecular tests1,2. In cases where there is enough tissue from a biopsy for molecular profiling, patients may have to wait several weeks or longer for results, or begin treatment without molecular results3,4. However, it is critical that informative molecular diagnosis be available given the advent of multiple targeted treatment options for patients diagnosed with NSCLC. Testing of circulating cell-free DNA (cfDNA) from liquid biopsy is a solution to the challenges of traditional tissue testing4,5,6. Current testing options for actionable mutations in NSCLC using cfDNA and a similar dPCR-based workflow for rapid result generation, include the epidermal growth factor receptor (EGFR) sensitizing mutations &#916;E746-A750 and L858R, EGFR resistance mutation T790M, KRAS Proto-Oncogene (KRAS) variants, and B-Raf Proto-Oncogene (BRAF) variant V600E. Although not as broadly adopted by the field, circulating tumor-derived messenger RNA (mRNA) isolated from liquid biopsy can also provide important clinical information7,8,9. We have previously developed and reported on methods of multiplexed detection of the Echinoderm Microtubule Associated Protein Like 4-Anaplastic Lymphoma Receptor Tyrosine Kinase (EML4-ALK) fusion variants from blood plasma10. In this study, we extended these methods to include higher-order multiplexed RNA targets for ROS1 and RET, covering eight fusion variants within each assay. The objective was to develop a rapid, sensitive, specific, and reproducible technique for the detection of these fusion variants from the plasma of patients previously diagnosed with NSCLC.
The test process is initiated in a physician&#39;s office using RNA stabilizing blood collection tubes11. These tubes contain a cell preservative as well as RNase inhibitors. Samples are shipped priority overnight to the centralized College of American Pathologists (CAP)-accredited/Clinical Laboratory Improvement Amendments (CLIA)-certified laboratory (Clinical Laboratory) for processing by competent personnel. Once received by the Clinical Laboratory, each step of processing is conducted under approved Standard Operating Procedures (SOP). Whole blood is centrifuged to recover plasma, which is then used to isolate circulating RNA that is either free in the blood or within encapsulating moieties, such as exosomes and platelets7,8,9. To isolate RNA from these compartments, we selected the system for RNA recovery based on comparisons of several extraction methods. The isolated RNA is concentrated and reverse transcribed to cDNA. Several reverse transcriptase enzymes and gene-specific primers were evaluated during the optimization of the cDNA synthesis method to maximize ROS1 and RET target transcript conversion10. This is critical for low abundance circulating transcripts, such as tumor-derived fusion variants. Finally, we optimized dPCR primer and probe concentrations to allow for multiplexed detection of RET or ROS1 fusion variants and the control gene, glucuronidase-&#946; (GUSB). We then combined the best conditions from each of the optimization studies into a final locked protocol before performing the analytic validation studies described in this report. This protocol and these results provide the basis for a rapid and sensitive workflow for the routine detection of rare fusion variants in circulation.

a blood-based test for the detection of ros1 and ret fusion transcripts from circulating ribonucleic acid using digital polymerase chain reaction

We have developed novel methods for the isolation and characterization of tumor-derived circulating ribonucleic acid (cRNA) for blood-based liquid biopsy. Robust detection of cRNA recovered from blood represents a solution to a critical unmet need in clinical diagnostics. The test begins with the collection of whole blood into blood collection tubes containing preservatives that stabilize cRNA. Cell-free, exosomal, and platelet-associated RNA is isolated from plasma in this test system. The cRNA is reverse transcribed to complementary DNA (cDNA) and amplified using digital polymerase chain reaction (dPCR). Samples are evaluated for both the target biomarker as well as a control gene. Test validation included limit of detection, accuracy, and robustness studies with analytic samples. The method developed as a result of these studies reproducibly detect multiple fusion variants for ROS1 (C-Ros proto-oncogene 1; 8 variants) and RET (rearranged during transfection proto-oncogene; 8 variants). The sample processing workflow has been optimized so that test results can consistently be generated within 72 hours of sample receipt.

This protocol describes a test system developed for the detection of RNA fusion variants for use in the measurement of driver mutations within the plasma of NSCLC patients (Figure 1A). Fusion mRNA products from the expression of the most common RET and ROS1 rearrangements in the NSCLC population were identified13,14,15,16,17. Multiplexed PCR assays were then designed to detect the eight most common transcript variants for each target in NSCLC within a single reaction. The most common translocations at the ROS1 locus generate associations with the 5&#39; portions of the CD74, SDC4, SLC34A2, EZR or TPM3 genes (Figure 1B). The most common translocations at the RET locus lead to juxtaposition with KIF5B, for which the assay covers six exon junctions. Additional RET partners that are covered include those with CCDC6 and TRIM33 (Figure 1C). In total, the assays cover approximately 88% of ROS1 and 99% of RET alterations known to occur in the NSCLC patient population17.
The specificity of the assay components was first evaluated using eight individual in vitro RNAs that contain the mRNA sequence for the fusion transcripts covered by the ROS1 or RET multiplexed assays. Each RNA species was tested against each individual variant assay that comprises the multiplexed version. There was no cross-reactivity of these assays, thus demonstrating 100% analytic specificity within the designed multiplexed assays (data not shown). To determine the lower limit of detection of the test protocol, total RNA derived from cell lines expressing a fusion variant included in the assay were mixed into a background of normal RNA at 5%, 1%, 0.2%, and 0.04% concentrations. The multiplexed RET and ROS1 variant PCR assays detected as little as 0.2% fusion variant (Figure 2A-B). Additionally, a preparation of 5% off-target cell line derived RNA (expressing an EML4-ALK fusion transcript) was not detected with the multiplexed ROS1 and RET assays, further demonstrating specificity (Figure 2A-B).
Precision testing of the RT-dPCR process was performed for both ROS1 and RET. Analytic control material comprised of equimolar in vitro RNAs was processed at three concentrations (High, Medium, and Low) through reverse transcription and dPCR on three different occasions within the same day (intra-day), on three consecutive days (inter-day), and with two operators (inter-operator). Results from precision testing demonstrated accurate detection of both the fusion transcript of interest, as well as a control gene, GUSB, which is included as an internal QC metric (Figure 2C-D).
In addition to the GUSB internal control, each batch of clinical samples was run with a set of batch controls. A positive control was developed from a mixture of analytic in vitro RNA that represented each of the fusion variants tested in RT-dPCR, as well as analytic in vitro RNA for GUSB. This RNA was spiked into normal human plasma lysate during RNA extraction and was processed alongside the clinical samples throughout the protocol. The no reverse transcriptase (No RT) control was a negative control to confirm the absence of contaminating material in the RNA extraction workflow and demonstrate the specificity of the primers for RNA. The No RT control was generated using the same material as the positive control, but it does not include enzyme within the cDNA synthesis reaction. The no RNA control (NRC) is a negative control to confirm the absence of contaminating transcripts in the reverse transcription reaction components. This control was introduced into the workflow at the cDNA synthesis step, and water was added in the reaction instead of an RNA template. The No RT and NRC controls must be negative in both channels, if accurate results are to be delivered. Table 1 lists the reverse transcription reaction components for each control. Examples of the 2D plots for each of these controls are shown for the ROS1 (Figure 3 A-C) and RET (Figure 3 E-G) multiplex assays. Fusion variants were detected using a fluorescein amidite (FAM) probe and are represented along the y-axis, while the control gene, GUSB, was detected using a 5&#39;-hexachloro-fluorescein-CE phosphoramidite (HEX) probe and is on the x-axis. These batch controls were assessed over the course of 21 days to determine assay robustness. Fusion positive droplets and GUSB control gene droplets were observed for ROS1 and RET in all 21 runs executed over the course of the study (Figure 3D,H). All negative controls (No RT and NRC) yielded negative results across the entire 21 days (data not shown).
The ability to troubleshoot is a critical component of any test protocol to be run in the clinical laboratory setting. Here, we provide real world examples of sub-optimal results using the RT-dPCR protocol. The first is an example 2D plot demonstrating the importance of the no reverse transcriptase control (Figure 4A). In this example, mutant positive droplets were present even though there was no cDNA conversion due to lack of enzyme. This outcome was likely due to dPCR primers amplifying off-target genomic DNA. In this case, design of an intron-spanning assay will prevent amplification of genomic DNA. Alternatively, an RNase-free DNase enzyme can be used to eliminate the contaminating DNA, but this is not recommended for detection of rare targets, as some RNA degradation may occur during incubation with the enzyme. The next example 2D plot was a NRC with positive droplets in both channels (Figure 4B). This indicated contamination at some point in the RT-dPCR setup. In this case, the recommendation is to discard any potentially contaminated reagents used in testing, thoroughly decontaminate all equipment, and re-test with fresh reaction components. The third example 2D plot presented as a spray of droplets along a 45&#176; line (Figure 4C). This is often caused by shearing and coalescing of droplets. Careful droplet handling prior to thermal cycling is essential, as droplets are prone to damage. We recommend the use of automated droplet generation, when available. If transferring manually generated droplets, be certain to choose the recommended wide-bore tips and employ careful pipetting technique. Droplet transfer requires slow aspiration and dispensing, with each taking place over 5-6 seconds, and it is essential that the pipette tip opening does not touch the droplet cartridge or well. When dispensing, keep the pipette tip at the liquid level and raise it slowly as droplets are dispensed (view video for demonstration). The final 2D plot example demonstrates a lack of separation between the positive and negative droplet populations (Figure 4D). This can have several causes. Strong PCR inhibitors, such as detergents used in lysis buffers and an excess of highly degraded DNA, can cause loss of separation. In this case, consider adding a clean-up step between cDNA synthesis and dPCR (such as described within Step 5 of this protocol). Finally, lack of separation can also be due to sub-optimal amplification conditions, and optimization of the PCR step should also be considered.
Data within Figure 5 represent 984 real world patient sample turn-around times and demonstrates the rapid nature of this test workflow. Results were reported to the treating physician as early as within 48 hours (79% of cases) of sample receipt and in 95% of cases, within 72 hours. In conclusion, the use of stabilized circulating RNA blood collection tubes, optimized RNA extraction procedures from blood, and RT-dPCR run according to an optimized protocol with the appropriate internal and batch controls, can provide a rapid test system for the accurate detection of fusion RNA variants relevant in NSCLC.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57079/57079fig1v2.jpg" /> 
Figure 1: Overview of Blood Sample Processing Steps for Fusion Variant Detection using Assays Specific to the Most Prevalent RET and ROS1 Variants in NSCLC. (A) Sample testing is initiated when whole blood is drawn and a BCT is shipped within the specimen collection kit to the Clinical Laboratory. Circulating RNA is recovered from multiple sources within the platelet-enriched plasma, reverse transcribed with gene specific priming, and purified for use in dPCR. Samples are processed using a commercially available system which consists of droplet generation (emulsion), amplification, and droplet counting. Data is analyzed using commercially available software. The test results are then documented and reported back to the test-requesting physician. The process is designed to work within a timeframe of 72 hours from sample receipt to result release. Eight variants for (B) ROS1 and (C) RET are covered within the multiplexed assays. Adapted from Biodesix website with permission. <a href="https://cloudflare.jove.com/files/ftp_upload/57079/57079fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57079/57079fig2.jpg" /> 
Figure 2: Analytic Validation. Cell-lines expressing (A) SDC4-ROS1 fusion and (B) CCDC6-RET fusion were diluted in a background of total human wild-type RNA (WT RNA). With each fusion variant, the limit of detection was established at 0.2% variant frequency using pre-defined criteria for each variant assay. All samples above this threshold also contained at least 21 copies of control gene. 5% EML4-ALK (ALK) standard in a background of wild-type RNA was tested to demonstrate assay specificity, which was confirmed by a negative result. Analytic multiplexed RNA standards were measured at high, medium, and low concentrations for (C) ROS1 and (D) RET. Precision was evaluated over three runs on the same day (Intra-day), three runs on three consecutive days (Inter-day), and with two independent operators (Inter-operator). The means of copy number&#160;and standard deviations are shown. Adapted from Biodesix website with permission. <a href="https://cloudflare.jove.com/files/ftp_upload/57079/57079fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57079/57079fig3.jpg" /> 
Figure 3: Batch Processing Control Examples and Robustness Data. 2D plot of the ROS1 multiplexed assay dPCR results for (A) positive control, (B) no reverse transcriptase control, and (C) no RNA template control. (D) Controls were run on 21 consecutive days (excluding weekends and holidays). Mean copy number&#160;+/- standard deviations for ROS1 positive control were 439 +/- 141. No reverse transcriptase and no template controls were also run on each day, and these were all negative (data not shown). 2D plot of the RET multiplexed assay dPCR results for (E) positive control, (F) no reverse transcriptase control and (G) no RNA template control. (H) Controls were run on 21 consecutive days (excluding weekends and holidays). Mean copies +/- standard deviations for RET positive control were 586 +/- 182. Not shown are no reverse transcriptase and no template controls that were also run on each day and were all negative. Adapted from Biodesix website with permission. <a href="https://cloudflare.jove.com/files/ftp_upload/57079/57079fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57079/57079fig4.jpg" /> 
Figure 4: Troubleshooting RT-dPCR. 2D plots representing sub-optimal dPCR results obtained when there is (A) contamination within the no reverse transcriptase control, (B) contamination within the no RNA control, (C) shearing and coalescing of droplets, and (D) poorly optimized PCR conditions or PCR inhibition. <a href="https://cloudflare.jove.com/files/ftp_upload/57079/57079fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57079/57079fig5.jpg" /> 
Figure 5: Turnaround time (TAT). TAT (in hours) was compiled for tests requesting an RNA variant (n = 984). Data excludes weekends, holidays, and samples held for &#62;24 h due to incomplete clinical information on the laboratory Test Request Forms. <a href="https://cloudflare.jove.com/files/ftp_upload/57079/57079fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Table 1" src="/files/ftp_upload/57079/57079table1.jpg" /> 
Table 1: Preparation of Reverse Transcription Reagents for Process Controls.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component</td>
			<td>Volume</td>
		</tr>
		<tr>
			<td>2x dPCR supermix for probes 
			(no 2&#8217;-deoxyuridine 5&#8217;-triphosphate)</td>
			<td>10 &#181;L</td>
		</tr>
		<tr>
			<td>20x variant target primers/probes set 
			&#160;(450 nmol/L primers, 250 nmol/L FAM probe)</td>
			<td>1 &#181;L</td>
		</tr>
		<tr>
			<td>20x control target primers/probe set 
			(450 nmol/L primers, 250 nmol/L HEX probe)</td>
			<td>1 &#181;L</td>
		</tr>
		<tr>
			<td>nuclease-free water</td>
			<td>1 &#181;L</td>
		</tr>
		<tr>
			<td>cDNA</td>
			<td>7 &#181;L</td>
		</tr>
	</tbody>
</table>
Table 2: Preparation of the master mix for dPCR.
<table border="1" fo:display-align="center" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Cycling Step</td>
			<td>Temperature</td>
			<td>Time</td>
			<td># Cycles</td>
			<td>Ramp Rate</td>
		</tr>
		<tr>
			<td>Enzyme activation</td>
			<td>95 &#176;C</td>
			<td>10 min</td>
			<td>1</td>
			<td rowspan="4">~2 oC/s</td>
		</tr>
		<tr>
			<td>Denaturation</td>
			<td>94 &#176;C</td>
			<td>30 s</td>
			<td rowspan="2">40</td>
		</tr>
		<tr>
			<td>Annealing/extension</td>
			<td>55 &#176;C</td>
			<td>1 min</td>
		</tr>
		<tr>
			<td>Enzyme deactivation</td>
			<td>98 &#176;C</td>
			<td>10 min</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Hold (optional)</td>
			<td>4 &#176;C</td>
			<td>infinite</td>
			<td>1</td>
			<td>~1 oC/s</td>
		</tr>
	</tbody>
</table>
Table 3: Thermal Cycling Conditions.

Watch this Scientific Journal Video about A Blood-based Test for the Detection of ROS1 and RET Fusion Transcripts from Circulating Ribonucleic Acid Using Digital Polymerase Chain Reaction at JoVE.com

A Blood-based Test for the Detection of ROS1 and RET Fusion Transcripts from Circulating Ribonucleic Acid Using Digital Polymerase Chain Reaction

Detection of circulating ribonucleic acid (cRNA) from blood is an unmet need in clinical diagnostics. Here we describe methods that characterize cRNA from non-small cell lung cancer patients using sensitive and specific digital polymerase chain reaction. The tests meet design requirements to detect fusion variants within 72 hours.

We have developed novel methods for the isolation and characterization of tumor-derived circulating ribonucleic acid (cRNA) for blood-based liquid biopsy. ...

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10.2K Views.  Biodesix. Detection of circulating ribonucleic acid (cRNA) from blood is an unmet need in clinical diagnostics. Here we describe methods that characterize cRNA from non-small cell lung cancer patients using sensitive and specific digital polymerase chain reaction. The tests meet design requirements to detect fusion variants within 72 hours.

Video: A Blood-based Test for the Detection of ROS1 and RET Fusion Transcripts from Circulating Ribonucleic Acid Using Digital Polymerase Chain Reaction

The Trier Social Stress Test Protocol for Inducing Psychological Stress

This article demonstrates a psychological stress protocol for use in a laboratory setting. Protocols that allow researchers to study the biological pathways of the stress response in health and disease are fundamental to the progress of research in stress and anxiety.1 Although numerous protocols exist for inducing stress response in the laboratory, many neglect to provide a naturalistic context or to incorporate aspects of social and psychological stress. Of psychological stress protocols, meta-analysis suggests that the Trier Social Stress Test (TSST) is the most useful and appropriate standardized protocol for studies of stress hormone reactivity.2 In the original description of the TSST, researchers sought to design and evaluate a procedure capable of inducing a reliable stress response in the majority of healthy volunteers.3 These researchers found elevations in heart rate, blood pressure and several endocrine stress markers in response to the TSST (a psychological stressor) compared to a saline injection (a physical stressor).3 Although the TSST has been modified to meet the needs of various research groups, it generally consists of a waiting period upon arrival, anticipatory speech preparation, speech performance, and verbal arithmetic performance periods, followed by one or more recovery periods. The TSST requires participants to prepare and deliver a speech, and verbally respond to a challenging arithmetic problem in the presence of a socially evaluative audience.3 Social evaluation and uncontrollability have been identified as key components of stress induction by the TSST.4 In use for over a decade, the goal of the TSST is to systematically induce a stress response in order to measure differences in reactivity, anxiety and activation of the hypothalamic-pituitary-adrenal (HPA) or sympathetic-adrenal-medullary (SAM) axis during the task.1 Researchers generally assess changes in self-reported anxiety, physiological measures (e.g. heart rate), and/or neuroendocrine indices (e.g. the stress hormone cortisol) in response to the TSST. Many investigators have adopted salivary sampling for stress markers such as cortisol and alpha-amylase (a marker of autonomic nervous system activation) as an alternative to blood sampling to reduce the confounding stress of blood-collection techniques. In addition to changes experienced by an individual completing the TSST, researchers can compare changes between different treatment groups (e.g. clinical versus healthy control samples) or the effectiveness of stress-reducing interventions.1

This article describes a protocol for inducing psychological stress in participants, which enables researchers to measure psychological, physiological and neuroendocrine responses to stress within single participants or between groups.

This article demonstrates a psychological stress protocol for use in a laboratory setting.  Protocols that allow researchers to study the biological ...

The &alpha;-test: Rapid Cell-free CD4 Enumeration Using Whole Saliva

There is an urgent need for affordable CD4 enumeration to monitor HIV disease. CD4 enumeration is out of reach in resource-limited regions due to the time and temperature restrictions, technical sophistication, and cost of reagents, in particular monoclonal antibodies to measure CD4 on blood cells, the only currently acceptable method. A commonly used cost-saving and time-saving laboratory strategy is to calculate, rather than measure certain blood values. For example, LDL levels are calculated using the measured levels of total cholesterol, HDL, and triglycerides1. Thus, identification of cell-free correlates that directly regulate the number of CD4+ T cells could provide an accurate method for calculating CD4 counts due to the physiological relevance of the correlates.
The number of stem cells that enter blood and are destined to become circulating CD4+ T cells is determined by the chemokine CXCL12 and its receptor CXCR4 due to their influence on locomotion2. The process of stem cell locomotion into blood is additionally regulated by cell surface human leukocyte elastase (HLECS) and the HLECS-reactive active &alpha;1proteinase inhibitor (&alpha;1PI, &alpha;1antitrypsin, SerpinA1)3. In HIV-1 disease, &alpha;1PI is inactivated due to disease processes 4. In the early asymptomatic categories of HIV-1 disease, active &alpha;1PI was found to be below normal in 100% of untreated HIV-1 patients (median=12 &mu;M, and to achieve normal levels during the symptomatic categories4, 5. This pattern has been attributed to immune inactivation, not to insufficient synthesis, proteolytic inactivation, or oxygenation. We observed that in HIV-1 subjects with &gt;220 CD4 cells/&mu;l, CD4 counts were correlated with serum levels of active &alpha;1PI (r2=0.93, p&lt;0.0001, n=26) and inactive &alpha;1PI (r2=0.91, p&lt;0.0001, n=26) 5. Administration of &alpha;1PI to HIV-1 infected and uninfected subjects resulted in dramatic increases in CD4 counts suggesting &alpha;1PI participates in regulating the number of CD4+ T cells in blood 3.
With stimulation, whole saliva contains sufficient serous exudate (plasma containing proteinaceous material that passes through blood vessel walls into saliva) to allow measurement of active &alpha;1PI and the correlation of this measurement is evidence that it is an accurate method for calculating CD4 counts. Briefly, sialogogues such as chewing gum or citric acid stimulate the exudation of serum into whole mouth saliva. After stimulating serum exudation, the activity of serum &alpha;1PI in saliva is measured by its capacity to inhibit elastase activity. Porcine pancreatic elastase (PPE) is a readily available inexpensive source of elastase. PPE binds to &alpha;1PI forming a one-to-one complex that prevents PPE from cleaving its specific substrates, one of which is the colorimetric peptide, succinyl-L-Ala-L-Ala-L-Ala-p-nitroanilide (SA3NA). Incubating saliva with a saturating concentration of PPE for 10 min at room temperature allows the binding of PPE to all the active &alpha;1PI in saliva. The resulting inhibition of PPE by active &alpha;1PI can be measured by adding the PPE substrate SA3NA. (Figure 1). Although CD4 counts are measured in terms of blood volume (CD4 cells/&mu;l), the concentration of &alpha;1PI in saliva is related to the concentration of serum in saliva, not to volume of saliva since volume can vary considerably during the day and person to person6. However, virtually all the protein in saliva is due to serum content, and the protein content of saliva is measurable7. Thus, active &alpha;1PI in saliva is calculated as a ratio to saliva protein content and is termed the &alpha;1PI Index. Results presented herein demonstrate that the &alpha;1PI Index provides an accurate and precise physiologic method for calculating CD4 counts.

The &#945;-test: Rapid Cell-free CD4 Enumeration Using Whole Saliva

A CD4 enumeration method, the &#x03B1;-test, is described which uses whole saliva to provide rapid and accurate CD4 counts. The &alpha;-test costs pennies and eliminates the need for technical training, costly reagents such as monoclonal antibodies, instrumentation, refrigeration, transport of samples, as well as collection and handling of blood.

There is an urgent need for affordable CD4 enumeration to monitor HIV disease. CD4 enumeration is out of reach in resource-limited regions due to the time ...

Intraoperative Detection of Subtle Endometriosis: A Novel Paradigm for Detection and Treatment of Pelvic Pain Associated with the Loss of Peritoneal Integrity

Endometriosis is a common disease affecting 40 to 70% of reproductive-aged women with chronic pelvic pain (CPP) and/or infertility. The purpose of this study was to demonstrate the use of a blue dye (methylene blue) to stain peritoneal surfaces during laparoscopy (L/S) to detect the loss of peritoneal integrity in patients with pelvic pain and suspected endometriosis. Forty women with CPP and 5 women without pain were evaluated in this pilot study. During L/S, concentrated dye was sprayed onto peritoneal surfaces, then aspirated and rinsed with Lactated Ringers solution. Areas of localized dye uptake were evaluated for the presence of visible endometriotic lesions. Areas of intense peritoneal staining were resected and some fixed in 2.5% buffered gluteraldehyde and examined by scanning (SEM) electron microscopy. Blue dye uptake was more common in women with endometriosis and chronic pelvic pain than controls (85% vs. 40%). Resection of the blue stained areas revealed endometriosis by SEM and loss of peritoneal cell-cell contact compared to normal, non-staining peritoneum. Affected peritoneum was associated with visible endometriotic implants in most but not all patients. Subjective pain relief was reported in 80% of subjects. Based on scanning electron microscopy, we conclude that endometrial cells extend well beyond visible implants of endometriosis and appear to disrupt the underlying mesothelium. Subtle lesions of endometriosis could therefore cause pelvic pain by disruption of peritoneal integrity, allowing menstrual or ovulatory blood and associated pain factors access to underlying sensory nerves. Complete resection of affected peritoneum may provide a better long-term treatment for endometriosis and CPP. This simple technique appears to improve detection of subtle or near invisible endometriosis in women with CPP and minimal visual findings at L/S and may serve to elevate diagnostic accuracy for endometriosis at laparoscopy.

Loss of peritoneal integrity provides a new paradigm to understand and treat chronic pelvic pain in women with mild forms of endometriosis and can be easily detected using intraoperative instillation of dye at the time of laparoscopy.

Endometriosis is a  common disease affecting 40 to 70% of reproductive-aged women with chronic  pelvic pain (CPP) and/or infertility. The purpose of this ...

Cerenkov Luminescence Imaging (CLI) for Cancer Therapy Monitoring

In molecular imaging, positron emission tomography (PET) and optical imaging (OI) are two of the most important and thus most widely used modalities1-3. PET is characterized by its excellent sensitivity and quantification ability while OI is notable for non-radiation, relative low cost, short scanning time, high throughput, and wide availability to basic researchers. However, both modalities have their shortcomings as well. PET suffers from poor spatial resolution and high cost, while OI is mostly limited to preclinical applications because of its limited tissue penetration along with prominent scattering optical signals through the thickness of living tissues.
 Recently a bridge between PET and OI has emerged with the discovery of Cerenkov Luminescence Imaging (CLI)4-6. CLI is a new imaging modality that harnesses Cerenkov Radiation (CR) to image radionuclides with OI instruments. Russian Nobel laureate Alekseyevich Cerenkov and his colleagues originally discovered CR in 1934. It is a form of electromagnetic radiation emitted when a charged particle travels at a superluminal speed in a dielectric medium7,8. The charged particle, whether positron or electron, perturbs the electromagnetic field of the medium by displacing the electrons in its atoms. After passing of the disruption photons are emitted as the displaced electrons return to the ground state. For instance, one 18F decay was estimated to produce an average of 3 photons in water5. 
 Since its emergence, CLI has been investigated for its use in a variety of preclinical applications including in vivo tumor imaging, reporter gene imaging, radiotracer development, multimodality imaging, among others4,5,9,10,11. The most important reason why CLI has enjoyed much success so far is that this new technology takes advantage of the low cost and wide availability of OI to image radionuclides, which used to be imaged only by more expensive and less available nuclear imaging modalities such as PET.
 Here, we present the method of using CLI to monitor cancer drug therapy. Our group has recently investigated this new application and validated its feasibility by a proof-of-concept study12. We demonstrated that CLI and PET exhibited excellent correlations across different tumor xenografts and imaging probes. This is consistent with the overarching principle of CR that CLI essentially visualizes the same radionuclides as PET. We selected Bevacizumab (Avastin; Genentech/Roche) as our therapeutic agent because it is a well-known angiogenesis inhibitor13,14. Maturation of this technology in the near future can be envisioned to have a significant impact on preclinical drug development, screening, as well as therapy monitoring of patients receiving treatments.

Cerenkov Luminescence Imaging CLI for Cancer Therapy Monitoring

Use of Cerenkov Luminescence Imaging (CLI) for monitoring preclinical cancer treatment is described here. This method takes advantage of Cerenkov Radiation (CR) and optical imaging (OI) to visualize radiolabeled probes and thus provides an alternative to PET in preclinical therapeutic monitoring and drug screening.

In  molecular imaging, positron emission tomography (PET) and optical imaging (OI)  are two of the most important and thus most widely used modalities1-3. ...

Assessment of Gastric Emptying in Non-obese Diabetic Mice Using a [13C]-octanoic Acid Breath Test

Gastric emptying studies in mice have been limited by the inability to follow gastric emptying changes in the same animal since the most commonly used techniques require killing of the animals and postmortem recovery of the meal1,2. This approach prevents longitudinal studies to determine changes in gastric emptying with age and progression of disease. The commonly used [13C]-octanoic acid breath test for humans3 has been modified for use in mice4-6 and rats7 and we previously showed that this test is reliable and responsive to changes in gastric emptying in response to drugs and during diabetic disease progression8. In this video presentation the principle and practical implementation of this modified test is explained. As in the previous study, NOD LtJ mice are used, a model of type 1 diabetes9. A proportion of these mice develop the symptoms of gastroparesis, a complication of diabetes characterized by delayed gastric emptying without mechanical obstruction of the stomach10.
This paper demonstrates how to train the mice for testing, how to prepare the test meal and obtain 4 hr gastric emptying data and how to analyze the obtained data. The carbon isotope analyzer used in the present study is suitable for the automatic sampling of the air samples from up to 12 mice at the same time. This technique allows the longitudinal follow-up of gastric emptying from larger groups of mice with diabetes or other long-standing diseases.

Assessment of Gastric Emptying in Non-obese Diabetic Mice Using a [13C]-octanoic Acid Breath Test

Determination of gastric emptying with a non-invasive [13C]-octanoic acid breath test for tracking gastroparesis in female NOD LtJ mice.

Gastric emptying studies in mice have been limited by the inability to follow gastric emptying changes in the same animal since the most commonly used ...

Isolation and Propagation of Circulating Tumor Cells from a Mouse Cancer Model

Cancer metastasis is the foremost cause of cancer-associated deaths. Recent studies have shown that circulating tumor cells (CTCs) are important in cancer metastasis. Indeed, the number of CTCs correlates with tumor size. Here, a detailed description is provided of a methodology for isolation and propagation of CTCs from a syngeneic mouse model of hepatocellular carcinoma (HCC) which allows for downstream analysis of potentially important molecular mechanisms of solid organ tumor metastasis. This method is efficient and reproducible. It is a non-invasive technique and, therefore, has potential to replace the invasive biopsy of tissues from humans which may be associated with complications. Therefore, the method discussed here allows for the isolation and propagation of CTCs from whole blood samples such that they can be examined and characterized. This has potential for future adaptation for clinical applications such as diagnosis, and personalized targeted therapy.

Circulating tumor cells (CTCs) have been shown to play an important role in tumor metastasis. Here, a method for the isolation and propagation of CTCs from the whole blood of a syngeneic mouse tumor model of hepatocellular carcinoma (HCC) metastasis is described.

Cancer metastasis is the foremost cause of cancer-associated deaths. Recent studies have shown that circulating tumor cells (CTCs) are important in cancer ...

Novel Diagnostics in Revision Arthroplasty: Implant Sonication and Multiplex Polymerase Chain Reaction

In orthopedic patients, foreign body-associated infections, especially periprosthetic joint infections (PJIs), are a devastating complication of arthroplasty. Infection requires complex treatment, may result in long hospitalization and causes considerable costs. Multiple surgical revisions can be necessary in these patients, with a loss in function as well as in quality of life.
The routine preoperative diagnostics include blood examination for C-reactive protein (CRP) and other biomarkers, as well as joint aspirate analysis for cell count, differentiation, and culture. Intraoperative specimens for histology and microbiology are also standard procedure. The microbiological examination of removed implants with sonication, in combination with the implementation of molecular biology techniques in microbiology, represent two novel techniques currently employed to enhance the differential diagnostics of PJI.
We present here the step-wise procedure of analyzing joint aspirate and sonication fluid, using a cartridge-based multiplex polymerase chain reaction (PCR) system. Results were matched against conventional cultures and consensus criteria for PJI. Conventional microbiological cultures from tissue biopsies, joint aspirate and sonication fluid showed a sensitivity of 66.7%, 66.7%, and 88.9%, respectively, and a specificity of 82.3%, 54.6%, and 61.5%, respectively. The PCR diagnostic of the sonication fluid and the joint fluid showed a sensitivity of 50.0% and 55.6%, respectively, and both a specificity of 100.0%. Both PCR diagnostics combined had a sensitivity of 66.7% and a specificity of 100.0%. The multiplex PCR therefore presents a rapid diagnostic tool with moderate sensitivity but high specificity in diagnosing PJI.

Differential diagnostics in painful arthroplasty is crucial for treatment success. Joint aspiration is routinely performed preoperatively. Multiplex polymerase chain reaction of the joint aspirate and the sonication fluid, a feasible tool for quick pathogen detection from these samples is described.

In orthopedic patients, foreign body-associated infections, especially periprosthetic joint infections (PJIs), are a devastating complication of ...

Taste Exam: A Brief and Validated Test

The emerging importance of taste in medicine and biomedical research, and new knowledge about its genetic underpinnings, has motivated us to supplement classic taste-testing methods in two ways. First, we explain how to do a brief assessment of the mouth, including the tongue, to ensure that taste papillae are present and to note evidence of relevant disease. Second, we draw on genetics to validate taste test data by comparing reports of perceived bitterness intensity and inborn receptor genotypes. Discordance between objective measures of genotype and subjective reports of taste experience can identify data collection errors, distracted subjects or those who have not understood or followed instructions. Our expectation is that fast and valid taste tests may persuade researchers and clinicians to assess taste regularly, making taste testing as common as testing for hearing and vision. Finally, because many tissues of the body express taste receptors, taste responses may provide a proxy for tissue sensitivity elsewhere in the body and, thereby, serve as a rapid, point-of-care test to guide diagnosis and a research tool to evaluate taste receptor protein function.

This protocol measures human taste responses and includes a brief anatomical assessment, a short taste test, and a validation method using the subject&#39;s reported sensation and taste receptor genotype.

The emerging importance of taste in medicine and biomedical research, and new knowledge about its genetic underpinnings, has motivated us to supplement ...

Digital PCR for Quantifying Circulating MicroRNAs in Acute Myocardial Infarction and Cardiovascular Disease

Circulating serum microRNAs (miRNAs) have shown promise as biomarkers for the cardiovascular disease and acute myocardial infarction (AMI), being released from the cardiovascular cells into the circulation. Circulating miRNAs are highly stable and can be quantified. The quantitative expression of specific miRNAs can be linked to the pathology, and some miRNAs show high tissue and disease specificity. Finding novel biomarkers for cardiovascular diseases is of importance for medical research. Quite recently, digital polymerase chain reaction (dPCR) has been invented. dPCR, combined with fluorescent hydrolysis probes, enables specific direct absolute quantification. dPCR exhibits superior technical qualities, including a low variability, high linearity, and high sensitivity compared to the quantitative polymerase chain reaction (qPCR). Thus, dPCR is a more accurate and reproducible method for directly quantifying miRNAs, particularly for the use in large multi-center cardiovascular clinical trials. In this publication, we describe how to effectively perform digital PCR in order to assess the absolute copy number in serum samples.

Circulating microRNAs have shown promise as biomarkers for cardiovascular diseases and acute myocardial infarctions. In this study, we describe a protocol for miRNA extraction, reverse transcription, and digital PCR for the absolute quantification of miRNAs in the serum of patients with cardiovascular disease.

Circulating serum microRNAs (miRNAs) have shown promise as biomarkers for the cardiovascular disease and acute myocardial infarction (AMI), being released ...

Traditional Trail Making Test Modified into Brand-new Assessment Tools: Digital and Walking Trail Making Test

The Trail Making Test (TMT) is a well-accepted tool for evaluating executive function. The standard TMT was invented more than 60 years ago and has been modified into many versions. With the development of digital technologies, TMT is now modified to a digitized version. The present study demonstrated digital TMT (dTMT) performed on a computer, and Walking TMT (WTMT) on the floor. Both revealed more information compared with the traditional version of TMT.

Here, we present a protocol to show how to perform two types of cognitive assessment tools derived from the paper-pencil version of the Trail Making Test.

The Trail Making Test (TMT) is a well-accepted tool for evaluating executive function. The standard TMT was invented more than 60 years ago and has been ...