Application Protocol: DNA quantification by using Absolute Quantitative PCR (qpcr)

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1 REPRODUCTION AND USE This document is protected by copyright and cannot be used or shared without permission from Vortex Biosciences, Inc. Such permission is given on condition that Vortex Biosciences is acknowledged whenever this protocol is reproduced or is published in whole or in part. PURPOSE This Application Protocol describes a SYBR green based qpcr method for quantifying DNA isolated from cells. These cells have been potentially enriched with Vortex technology from a human blood sample. SYBR Green is a dye that intercalates with double-stranded DNA. This intercalation causes the SYBR to fluoresce (see Figure 1, modified from The qpcr machine detects the fluorescence and the software calculates Ct values from the fluorescence intensity. This protocol is designed for a 7500 Fast Real-Time PCR machine from Applied Biosystems, the 7500 Software v2.02 and a SYBR Green from Roche (reference available in the next section). To use another PCR system or another type of SYBR Green, please refer to the vendor manual. Figure 1. SYBR Green qpcr Reaction scheme Page 1 of 10

2 MATERIALS Consumables Normal waste container. Paper towels. Filtered pipette tips (p2, p10, p20, p200, p1000). Aluminium Foil and Parafilm. Kimwipes. 1.5mL Maxymum Recovery microcentrifuge tubes or equivalent (Axygen, ref # MCT-150-L-C). MicroAmp optical adhesive film (Applied Biosystems, ref # ). 96 well qpcr plate (Applied Biosystems, ref # ). 8-Strip 0.2 ml PCR Tubes (GeneMate, ref # T ). Equipment Tube rack. Pipettors (p2, p10, p20, p200, p1000) and multichannel pipettor. Vortexer. Benchtop microcentrifuge. Benchtop Mini centrifuge. Centrifuge with buckets for well plates, Beckman GS-6R or equivalent. Base for qpcr plate (MicroAmp, Splash Free 96-Well Base, ref # ). Plate sealing paddle. Applied Biosystems 7500 Fast Real-Time PCR system or equivalent. AirClean 600 PCR Workstation or equivalent. Relevant Personal Protective Equipment (PPE). Kits and Reagents 10% Bleach. Ethanol 70%. ACROS Organics Ethanol, Absolute (200 Proof), ACS reagent or equivalent. UltraPure distilled water (DNase, RNase free) (Invitrogen, ref # ). Human genomic DNA (200 ng/µl) (Roche, ref # ). hline1 Forward and Reverse Primers (desalted, from IDT). (Forward: 5 -TCACTCAAAGCCGCTCAACTAC-3 Reverse: 5 -TCTGCCTTCATTTCGTTATGTACC-3. From: Rago C, et al. Cancer Res. 2007; 67: ). 2X SYBR green master mix (Roche, ref # ). Page 2 of 10

3 PROCEDURE Primer Preparation The primers only need to be prepared for the first use. a) Stock solution: Make each primer to a 100 µm stock solution by dissolving the primer powder into UltraPure water. Store the solutions at -20 C for up to 2 years. b) Working solution: Prepare the working primer mix so that it contains 10 µm of each primer. It is recommended to prepare a final volume of 200 µl. Store the solution at -20 C up to 6 months. Setting up the PCR hood Wipe the PCR hood (both surface and the interior walls) with 10% bleach first and then 70% ethanol. Place materials to be used for the following procedure inside the hood on the solid work surface. Materials taken into the cabinet should be kept to a minimum but should be good enough to avoid repeatedly bringing equipment or reagents in and out of the hood. Disinfect the exterior of these materials with 70% ethanol prior to bring inside the hood. Keep one set of dedicated pipettes inside the PCR hood if it is possible. Close the door and turn on the UV light of the PCR hood for 15 min. UV rays are dangerous to the eyes and skin when exposed without protection. Keep doors closed at all times until the UV light turns off. When the UV cycle is done, open the door, turn on the blower and set up the following experiment inside the PCR hood. Perform all work in the central area of the cabinet. Starting the experiment 1) Preparation of DNA standards a) Label 8 Maxymum Recovery microcentrifuge tubes as standards Standard 1 to Standard 8. b) Take the human genomic DNA (200ng/µL) from 4 C storage, flick the tube and briefly centrifuge with a benchtop Mini centrifuge. Keep on ice. c) Inside the PCR hood, prepare the standards 1 to 8 according to Table 1. First add water into the tube then add the DNA. Vortex and briefly centrifuge the tubes before pipetting for the next dilution. d) The standards can be stored at -20 C after each use with a maximum of 3 thaw and freeze cycles. Page 3 of 10

4 Table 1. Preparation of standards with serial dilutions Standard # DNA concentration (ng/µl) Volume to pipet (µl) From Volume of UltraPure water (µl) Standard Stock 184 Standard Standard Standard Standard Standard Standard Standard Standard Standard Standard Standard Standard Standard Standard ) Preparation of DNA samples It is not required to dilute the DNA extracted from rare cells such as CTC samples. If the DNA has been extracted from a large quantity of cells, such as a tissue biopsy or has been amplified with a Whole Genome Amplification (WGA) kit, it is recommended to measure the DNA concentration using a Qubit device, in order to have an approximation of the DNA concentration and determine if a dilution is needed (refer to DQ002). 3) Preparation of qpcr master mix a) Thaw the primer mix and the 2XSYBR green master mix from -20 C, flick the tubes and briefly centrifuge. Keep on ice. b) Calculate the total number of reactions: Count the total number of samples, including the 8 standards. Then multiply by 2 duplicates. Always include 2-3 extra reactions to allow for pipetting errors. c) Prepare the qpcr master mix in a Maxymum Recovery microcentrifuge tube according to the following table. Table 2. PCR reaction contents Content (ul) 1 reaction n reactions 2XSYBR Green master mix 10 10*n hline1 primer mix 1 1*n UltraPure water 7 7*n Subtotal 18 DNA template 2 Total 20 d) Vortex and briefly centrifuge the qpcr master mix. Keep on ice. 4) Preparation of the 96-well qpcr plate a) Label the qpcr plate accordingly to the samples referred in Table 3. Page 4 of 10

5 Table 3. Example of template of a 96-well qpcr plate with 10 samples in duplicate (Std: Standard, S: sample, NTC: Non Template Control) A Std 1 Std 1 S1 S1 S9 S9 B Std 2 Std 2 S2 S2 S10 S10 C Std 3 Std 3 S3 S3 NTC NTC D Std 4 Std 4 S4 S4 E Std 5 Std 5 S5 S5 F Std 6 Std 6 S6 S6 G Std 7 Std 7 S7 S7 H Std 8 Std 8 S8 S8 b) Aliquot 18 µl of the qpcr master mix in each reaction well. c) Add 2 µl of each DNA standard, DNA Samples to the corresponding wells. For the NTC, use 2 µl of UltraPure water instead. d) Carefully seal the plate with a MicroAmp optical adhesive film by using a sealing paddle. Note: Seal the plate tightly and nicely to avoid possible evaporation of the samples and result bias. You may also choose not to use the most external wells such as wells in row A, H and wells in column 1 and 12. e) Centrifuge the PCR plate up to 250 g (1000 rpm) for 30 seconds. f) Perform the qpcr. You may wrap the plate in aluminum foil to avoid direct light and put it at 4 C during the time when you set up the qpcr instrument. 5) Perform qpcr on 7500 qpcr instrument. a) Please refer to the APPENDIX or 7500 qpcr menu to set up the PCR program. b) Perform PCR according to the PCR cycling condition listed in table 4. Table 4. PCR cycling condition Step Temp. Duration Hold 50 C 2 min Denature 95 C 10 min Cycle: 40x 95 C 15 sec 60 C 1 min Melt curve Note: This PCR cycling condition is based on the SYBR green mix from Roche (ref # ). This cycling condition needs to be adjusted accordingly based on the SYBR Green from different vendors. 6) Data analysis to determine the amount of DNA in each sample. Get the data from the qpcr instrument by exporting the file to an excel format file (refer to APPENDIX) and then do data analysis (refer to the qpcr DATA ANALYSIS section). Page 5 of 10

6 Clean up Bring all equipment and supplies back once the experiment is done. Spray & wipe down all surfaces with 10% bleach and 70% ethanol. Keep the shared space clean for next users. Throw away the well-plate after each PCR run is completed. qpcr Data analysis 1) Establish the standard curve The standard curve is plotted by using the Ct (cycle threshold) values of standards and the corresponding Log (DNA concentration). Refer to Figure 2. The qpcr software can generate the standard curve automatically as long as the template of the experiment was set up appropriately. You may also plot your own standard curve in the excel file if you want to double check it. The R-squared value (coefficient of determination) is a critical parameter for the evaluation of the PCR efficiency. When the R-squared value is 1, the value of Y (Ct) can be used to accurately calculate the value of X (Log DNA conc.). A R-squared value > 0.99 provides good confidence in correlating Y (Ct) and X (Log Concentration) values. Ideally the efficiency of a PCR should be 100%, meaning that for each cycle the amount of product doubles. For an efficiency of 100%, the slope is A good reaction should have an efficiency between 90% and 110%, which corresponds to a slope between and Figure 2. Standard curve Page 6 of 10

7 2) Check the melting curve. It is important to run a dissociation curve following the qpcr for SYBR Green-based amplicon detection. This is due to the fact that SYBR Green will detect any double stranded DNA including primer dimers, contaminating DNA, and nonspecific PCR amplification. Viewing a dissociation curve ensures that the desired amplicon is detected. A typical plot of the derivative melting curve is depicted on Figure 3. A single peak indicates specific amplification of the desired target and absence of contaminating products in the reaction. Contaminating DNA or primer dimers would show up as an additional peak separated from the desired amplicon peak. If the additional peak(s) was found, please refer to the troubleshooting section. Figure 3. Derivative Melting Curve for Standards 3) Data analysis If the standard curve looks good (good R-squared value), use directly the concentration of DNA in ng/µl (column quantity ) indicated in the file. Otherwise, please refer to the troubleshooting section then repeat the experiment until a good standard curve is obtained. Since each sample was processed in duplicate, calculate the average of the DNA concentration in ng/µl for each sample then multiply by the factor of dilution (if a dilution of the sample was used). The final concentration of DNA (ng/µl) is thus obtained for each sample. Finally, multiply the final concentration of DNA by the DNA volume to get the total amount of DNA for each sample. DOWNSTREAM APPLICATIONS The qpcr quantified DNA could be used for PCR direct sequencing, multiplex PCR, whole genome amplification and NGS sequencing. Page 7 of 10

8 DOCUMENTATION It is recommended to keep track of the sample processing in a database and update it at the end of each DNA extraction round. [Protocol Vortex DA002. Whole genome amplification (WGA) by using REPLI-g kit]. This protocol explains how to do the WGA from a small amount of fresh cells or a small amount of DNA extracted from fresh cells. [Protocol Vortex DS001. PCR based Sanger sequencing]. This protocol explains how to detect a gene mutation by using PCR and Sanger sequencing. [Protocol Vortex DQ002. DNA quantification by Qubit 3.0 Fluorometer]. This protocol is used to determine DNA concentration by using Qubit 3.0 Fluorometer. TROUBLESHOOTING Problem Cause Solution NTC positive amplification Contamination. Primer dimers formation. Repeat the experiment with fresh clean water and fresh diluted primers. Clean the bench, pipettes etc. to eliminate contamination. No amplification of Standards Pipetting error / Reagents missing. SYBR master mix loses efficiency Calculate volumes for each step carefully. Add each reagent precisely and pipet accurately. Check the master mix for expiration or accidental storage at room temp for a long time. Sample concentration is out of the linear range of the standard curve. DNA sample s concentration is too high or too low. If the concentration is too high, dilute the DNA sample and repeat the experiment. If the concentration is too low, repeat the experiment with more DNA. Poor qpcr efficiency/slope Poor pipetting of identical replicates or poor pipetting of standards. The standards were made with wrong concentration. The standards were thawed and frozen more than three times. Repeat the experiment by pipetting accurately using well calibrated pipettes and appropriate tips. Repeat the experiment with new freshly prepared standards. Page 8 of 10

9 Problem Cause Solution High well-to-well variability Inaccurate pipetting. Evaporation. Repeat the experiment by pipetting accurately using well calibrated pipettes and appropriate tips. Seal the plate tightly. IMPORTANT NOTES 1. Avoid contamination of samples and DNA cross-contamination Clean the set up area and the set of pipettes with 10% bleach and 70% ethanol. Use the dedicated set of pipettes for PCR only and keep it inside the hood. Accuracy in the pipetting and thorough mixing is very important for the accuracy of the qpcr. Be careful to minimize pipetting errors and to mix each solution thoroughly. Always wear PPE (lab coat and gloves made of latex, nitrile or vinyl). Change gloves frequently. When possible, avoid opening tubes by hand. Use Kelly forceps previously disinfected with 70% ethanol instead. Be careful when working with tubes or wells close to each other. Process one sample at a time and close each tube right after it is finished to be processed, before starting with another sample. When working with a plate, be careful not to cross-contaminate the samples. 2. Keep track Always write down the date, the sample ID, the procedure and your initials on the tubes. Update DNA database, if available. REFERENCES Roche FastStart Universal SYBR Green Master (Rox) data sheet. Rago C, et al. Cancer Res. 2007; 67: Manual of Applied Biosystems 7500 Real-Time PCR Systems. Manual of AirClean 600 PCR Workstation. Page 9 of 10

10 APPENDIX: Use the 7500 qpcr machine for the experiment 1. Turn on the qpcr machine and the connected computer. 2. Open the 7500 Software. Log in as Guest. 3. Click on Advanced Set up. 4. Experiment Property setup. In the Setup section and the Experiment Properties subsection on the left of the screen, enter or select: 1) Identifying the experiment: Enter the experiment name. 2) Instrument used to run the experiment: Select 7500 fast (96 wells). 3) Type of experiment to setup: Select Quantitation-Standard Curve. 4) Reagent used to detect targeted sequence: Select SYBR green reagents. 5) Ramp speed to use in the instrument run: Select Standard (~ 2 hours to complete the run). 5. Plate setup. In the Setup section, click on the subsection Plate set up. 1) In the section Define targets & samples. 2) Enter the target name (hline1), and the reporter (SYBR green). 3) Enter all the samples. For each sample enter the date of processing and the sample ID. 4) Click on Assign targets & samples. 5) On the plate template, right click and select Define & setup standards. 6) Select the target for the standard (part Select a target ): eg. hline1. 7) Enter the following information (part Define the standard curve ): 7.1) Number of points: ) Number of replicates: 2 for the duplicate. 7.3) DNA quantity at the beginning: ) Serial factor: 1:4. 8) Click on Let me select wells (part Select and arrange wells for the standards ). 9) On the template, select the wells corresponding to the standards. 10) Tick the box Arrange the standards in rows. 11) Click on Apply, then Close. 12) Assign the samples and negative control: on the well plate template, select the wells corresponding to sample 1 and tick the box sample 1 in the part Assign sample to the selected wells. 13) Repeat step 12) for each sample. 14) Assign target to the selected wells. 15) Select on the template all wells corresponding to the samples, then click on U for Unknown in the part Assign target to the selected wells. 16) Select on the template the wells corresponding to the negative control, then click on N in the part Assign target to the selected wells. 17) Save the file. Use.eds format. 6. Run Method Setup. In the menu on the left, section Setup, click on the subsection Run Method. Then set the temperature, time and cycles based on PCR cycling condition listed in the table Load the plate in the qpcr machine and click on Start Run. 8. At the end of the run, check the Standard curve plot and the Melting curve. 9. Export the data to an excel format file: 1) Click on File then on Export. 2) Select data to export: Select Results 3) Select to export data in One File. 4) Enter the export file name or use the current name. 5) Choose the Export File Location. 6) Click on Start Export. 10. Proceed to the data analysis to determine the amount of DNA in each sample. Page 10 of 10