First-strand complementary DNA synthesis protocol: using Agilent MassCode complementary DNA synthesis kit. Cat# 5190–3553

Note: wear gloves at all times during the first-strand cDNA synthesis and PCR amplification procedures and while handling materials and equipment to prevent contamination by ribonucleases (RNases).

1. Prepare the required amount of master mix 1 with 10% overage for first-strand cDNA synthesis reaction:
   1. Note: mix each component and spin down before use.
   2. Master mix 1 (per sample).
   3. 3 µl of random primers (0.1 µg/µl).
   4. 1.7 µl of RNase-free water.
2. Prepare the cDNA synthesis reaction in a microcentrifuge tube:
   1. 5.7 µl of master mix 1 per microcentrifuge tube.
   2. 10 µl of nucleic acids isolated from clinical sample or positive control.
3. Incubate the reaction at 70 °C for 10 minutes.
4. Flash cool the reaction by transferring directly to ice.
5. Prepare the required amount of master mix 2 with 10% overage for first-strand cDNA synthesis reaction:
   1. Master mix 2 (per sample + 10% overage).
   2. 2.0 µl of 10 × MassCode Reverse Transcriptase Buffer.
   3. 0.8 µl of dNTP mix (25 mM of each dNTP).
   4. 0.5 µl of RNase Block Ribonuclease Inhibitor (40 units/µl).*
   5. 1 µl of MassCode Multiple Temperature Reverse Transcriptase.*
6. Add 4.3 µl of master mix 2 per reaction.*
   1. Note: To prevent heat inactivation, MassCode Reverse Transcriptase and RNase Block must be added after the 70 °C incubation is completed and the reaction has cooled.
7. Transfer tubes to thermal cycler and run on following cycle:
   1. 25 °C, 10 minutes, primer extension.
   2. 42 °C, 60 minutes, strand synthesis.
   3. 70 °C, 15 minutes, terminate reaction.

[*DO NOT VORTEX OR CENTRIFUGE. KEEP ON ICE]

Polymerase chain reaction protocol: using 2x MassCode polymerase chain reaction master mix. Cat# 5190–3745 and Multiplex Tagged Primer Mix

MassCode Tags are light sensitive. Minimise exposure to light as much as possible!

If using ready-made Primer mix go to step 6; or to make Primer mix:

1. Reconstitute each lyophilised primer with RNase/DNase-free water to make a final concentration of 100 µM.
2. Mix an equal volume of each primer to make a master mix. For example, to make a primer mix for 20-plex assay combine 100 µl of each primer × 40 primers = 4000 µl, then aliquot them 500 µl × 8 tubes (see Table 46).
3. The final concentration of each primer in PCR is 250 nM. You have to add 0.05 µl of each primer per reaction or 0.1 µl of primer set (two primers).
4. For example, for 20-plex assay you have to add 0.05 µl × 40 primers = 2 µl of primer mix (or 0.1 µl × 20-plex = 2 µl) per 1 reaction. For 100 reactions (1 plate) 2 µl × 100 = 200 µl of primer mix.
5. For example, for 25-plex assay (50 primers) add 0.1 × 25-plex= 2.5 µl/reaction or 250 µl per plate (100 reactions).
6. Prepare the required amount of PCR master mix with 10% overage and aliquot 16 µl into each PCR tube or 96-well plate well (Table 42).
7. Add 4 µl of the sample template, control template (provided), or water (for no template control) according to plate layout (Table 43).
8. Transfer to thermal cycler and run on following cycle (Table 44).

TABLE 46

Primer pairs within MassCode 14-plex panel

TABLE 42

PCR master mix for 13-plex (Big 11)

TABLE 43

Plate layout

TABLE 44

Cycles for MassCode PCR

Protocol for MassTag amplicon purification using a vacuum manifold. Cat# 5190–3745

Reminder: MassCode Tags are light sensitive. Minimise exposure to light as much as possible.

1. Prepare the DNA binding solution by adding an equal volume of 30% ethanol (final ethanol concentration is 15%). Put 20 ml per 96-well plate of DNA binding solution/ethanol mixture into a reservoir tray for a multichannel pipettor.
2. Using a multichannel pipettor, add 200 µl of DNA binding solution/ethanol mixture to each 20 µl PCR product in the PCR plate.
3. Place a MassCode 96-well binding plate on a vacuum manifold that contains a deep-well waste plate at the bottom. Using a multichannel pipette set for 250 µl, mix the contents of each well and then transfer the PCR product/DNA binding solution/ethanol mixtures into the wells of the binding plate.
   Note: If some of the 96 wells do not contain samples, seal the tops of the empty wells with tape.
4. Apply 400 mbar of vacuum to the binding plate until each well is dry. The vacuum force may decrease as the wells become dry (≈ 1 minute). Continue the vacuum for an additional 1 minute after the wells appear dry.
5. Release the vacuum and remove the binding plate from the manifold. Blot the bottom of the binding plate onto clean paper towels. Place the binding plate on top of a 96-well collection plate. Place a plate sealer on top of the binding plate. Tape the plate’s sides or use a rubber band to keep the two plates together to prevent an accidental spill. Centrifuge the plates at 1500 × g for 3 minutes at room temperature. Inspect the filter plate. Make sure the plate wells have been centrifuged until dry, repeat the centrifugation if there is any liquid still seen on the filters. Any buffer remaining will contribute to background noise in the mass spectrometer.
6. Wash #1. Add 500 µl of DNA Binding solution/ethanol mixture (from step #1) into each well of the DNA binding plate.
7. Apply 400 mbar of vacuum until each well is dry (≈ 1 minute). Continue the vacuum for an additional 1 minute after the wells appear dry.
8. Release the vacuum and remove the binding plate from the manifold. Blot the bottom of the binding plate onto clean paper towels. Place the binding plate on top of a 96-well collection plate. Place a plate sealer on top of the binding plate. Tape the plate’s sides or rubber band the two plates together to prevent an accidental spill. Centrifuge the plates at 1500 × g for 3 minutes at room temperature.
9. Open the vacuum manifold and discard the wash solution from the waste tray. Replace the waste tray inside and place the binding plate on top of the vacuum manifold.
10. Wash #2. Prepare 1 × PCR wash buffer by adding four volumes of 100% (v/v) ethanol to the 5 × PCR wash buffer container. After adding the ethanol, check the box on the label: [ □ ] 1 × (Ethanol Added). Store the 1 × PCR wash buffer tightly sealed at room temperature.
11. Add 700 µl of 1 × PCR wash buffer to each well of the binding plate.
12. Apply 400 mbar of vacuum until each well is dry (≈ 1 minute). Continue the vacuum for an additional 1 minute after the wells appear dry.
13. Release the vacuum and remove the binding plate from the manifold. Blot the bottom of the binding plate onto clean paper towels. Place the binding plate on top of a 96-well collection plate. Place a plate sealer on top of the binding plate. Tape the plate’s sides or rubber band the two plates together to prevent an accidental spill. Centrifuge the plates at 1500 × g for 3 minutes at room temperature.
14. Open the vacuum manifold and discard the wash solution from the waste tray. Replace the waste tray inside and place the binding plate on top of the vacuum manifold.
15. Wash #3. Prepare 80% ethanol wash solution by mixing 160 ml of 100% ethanol with 40 ml of nuclease-free water.
16. Add 700 µl of 80% ethanol wash solution to each well of the binding plate.
17. Apply 400 mbar of vacuum until each well is dry (≈ 1 minute). Continue the vacuum for an additional 1 minute after the wells appear dry.
18. Release the vacuum and remove the binding plate from the manifold. Blot the bottom of the binding plate onto clean paper towels. Place the binding plate on top of a 96-well collection plate. Place a plate sealer on top of the binding plate. Tape the plate’s sides or rubber band the two plates together to prevent an accidental spill. Centrifuge the plates at 1500 × g for 5 minutes at room temperature.
    Note: This step is to ensure that any ethanol from the wash solution is removed prior to adding elution buffer. Ethanol contamination in the eluted PCR product can cause quantification errors.
19. Remove the binding plate and 96-well collection plate from the centrifuge. Remove the 96-well collection plate and place the binding plate on top of a fresh 96-well collection plate that is suitable for the mass spectrometer autosampler (i.e. Agilent P/N 5042–1386). Make sure both plates are correctly oriented A1 to A1. Remove the plate sealer from the binding plate. Add 140 µl* of nuclease-free water directly onto the top of the fiber matrix at the bottom of each well using a multichannel pipette. Replace the plate sealer. Tape the plate’s sides or rubber band the two plates together to prevent an accidental spill.
20. Incubate the binding plate for 2 minutes at room temperature.
21. Centrifuge the binding plate and 96-well collection plate together at 1500 × g for 5 minutes at room temperature.
22. The purified MassCode amplicons are in the bottom of each well of the 96-well collection plate. Place a pre-slit well cap (Agilent P/N 5042–1389) on top of the 96-well collection plate for transfer to the mass spectrometer autosampler.

[*70 µl is the minimum volume required. Eluting in twice the volume (140 µl) allows for a duplicate mass spectrometric run of the samples.]

Protocol for mass spectrometer run

Ensure pre-run checks carried out and maintenance performed as detailed in MassCode maintenance guide.

1. Turn on UV unit – allow to warm up for 20–30 minutes until both lights are green.
2. Prepare mobile phase (1 l). Weigh out 0.46248 g liquid chromatography/MS-grade ammonium acetate (6 mM) and add to clean 1 l glass bottle. Add 500 ml liquid chromatography/MS-grade water and 500 ml liquid chromatography/MS-grade methanol. Swirl to mix.
3. Prepare system suitability calibrant. Defrost neat system suitability calibrant add 950 µl of liquid chromatography/MS-grade water and mix well. Transfer mixture to polypropylene vial suitable for MS autosampler.
4. Add 1 ml liquid chromatography/MS-grade water to glass vial suitable for MS autosampler.
5. Prepare run by placing water vial into vial slot 1 of autosampler (lid off), system suitability calibrant into vial slot 2 (lid off), and prepared 96-well plate into plate slot 1.
6. Set up plate details in MassCode software.
7. Check method set to ‘Enteric_13plex’ and run plate.

Target ions and calibrant mixes are detailed below and should be previously set up in ‘Target panel manager’ for ‘Enteric_13plex’.

Enteric 13-plex target panel as set up in ‘Target panel manager’ (Table 45).

TABLE 45

Target panel for MassCode assay

Nucleic acid extraction: using bead beating pre-step and QiaSymphony virus/pathogen kit. Cat#937055

Additional materials required:

• MS2 phage ssRNA – control for nucleic acids extraction and amplification (provided).
• STAR buffer (Roche; cat no 03335208001).
• Chloroform.
• Lysing matrix E (MPBiomedicals; cat no 116914500).

Note: MS2 is currently provided at two different concentrations:

1. 5 × 10⁵ copies MS2/µl.
2. 5 × 10⁶ copies/µl.

Amount of MS2 for extraction can vary depending on a sample type and the volume of the Elution Buffer recommended in the nucleic acids extraction protocol (50–200 µl).

We recommend using 5 × 10⁴ copies of MS2 for 50 µl of Elution Buffer, then 10 µl of extracted nucleic acids per RT cDNA synthesis reaction (10⁴ copies of MS2 per RT) and 4 µl of cDNA per 20 µl PCR. Based on our experience, 2 × 10³ copies of MS2 per PCR are optimal for detection of a good positive signal on MS. If a higher volume of Elution Buffer is required by an extraction protocol increase MS2 amount proportionally (e.g. 1 × 10⁵ copies of MS2 for 100 µl elution volume). Input amount of MS2 phage ssRNA can be increased or decreased depending on MassCode results.

Symphony loading

Waste Draw:

1. Affix waste bag to waste draw undercarriage.
2. Insert tip station, tip chute and waste vessels into waste draw ensuring that the tip station is fully inserted then close the waste draw.
3. On touchscreen select ‘scan now’ or ‘scan later’.
   Elution Draw:
4. Place black elution rack onto stainless carrier and place on slot 1 (cooled) in the elution draw.
5. On touchscreen select the rack and press ‘rack id’ button.
6. Scan the barcode on the Qiagen elution plate and place in elution rack in correct orientation and close the eluate draw.
7. On touchscreen press next and the Qiasymphony will briefly lock the draws and hood while scanning the elution plate. Once scanning is complete and the machine is unlocked, proceed to the next step.
   Reagents and Consumables Draw:
8. Remove the magnetic beads container from the reagents cassette and vortex for 3 minutes or until beads are evenly suspended and reinsert into reagent cassette.
9. Lift the reagent cartridges out of the white plastic stand and place into a grey cassette rack.
10. If the reagent cassette is being used for the first time then proceed to steps 11–15. If the cassette has been used before then go to steps 16–19.
11. Place a piercing lid on top of the cassette with the curved side of the lid on the opposite side of the cassette to the magnetic beads cartridge. Do not press down on the piercing lid or otherwise pierce the top of the reagent cartridges as the QiaSymphony will do this automatically.
12. Remove the seal from the magnetic beads trough ensuring that all parts of the seal are fully detached.
13. Using some clean tissue paper wipe away all droplets of the magnetic bead solution from around the rim of the bead container.
14. Slide the white enzyme rack onto the side of the reagents cassette and unscrew the proteinase K lids (the lids can be placed in the lid holders between the proteinase K rack and the cassette rack).
15. Proceed to step 20.
16. If the reagents cassette has been used before then remove the lids from the individual reagent cartridges and from the magnetic beads cartridge.
17. Using some clean tissue paper wipe away all droplets of the magnetic bead solution around the rim of the bead container.
18. Unscrew the proteinase K lids (the lids can be placed in the lid holders between the proteinase K rack and the cassette rack).
19. Proceed to step 20.
20. Load black 1500 µl and blue 200 µl tips, 8 rod covers and sample prep cartridges into reagents and consumables draw. For one run consisting of 24 samples the QIAsymphony will require 85 1500 µl tips, 24 200 µl tips, 3 8-rod covers and 18 sample prep cartridges. These quantities do not have to be exact but are the minimum amounts required.
21. On touchscreen press ‘bottle id’ button and scan ATL buffer bottle barcode and insert bottle into rear of reagents and consumables draw.
22. Load the reagents cassette that has been setup as described in steps 10–17.
23. Close the reagents and consumables draw and on touchscreen press ‘scan now’.
24. While the Qiasymphony is scanning, proceed to step 25.
    Samples Draw:
25. Make up carrier RNA in 2 ml microcentrifuge tubes suitable for the Qiasymphony using AVE buffer and carrier RNA stock. The amount required for any number of samples (up to 24) can be found in Table 47.
26. Unscrew the lids of the carrier RNA microcentrifuge tubes and place into a sample rack ensuring that the insert type in the rack matches the tube type. For 2 ml microcentrifuge tubes the insert has a red number above the barcode.
27. Unscrew the lids of the sample tubes and place them into a sample rack ensuring that the insert type in the rack matches the tube type. For 2 ml microcentrifuge tubes the insert has a red number above the barcode.
28. Only when the Qiasymphony has finished the scan will the sample drawer be unlocked. Once open, slide the samples rack up to the line in one of the first four slots and wait for the barcode scanner to align. Wait for the slot light to flash and then slide the rack past the barcode scanner until it clicks into place at the back of the draw. If correctly inserted the light will change from green to yellow. Do not place samples in the slot marked ‘A’ as this is for carrier RNA or internal controls only.
29. Slide the carrier RNA rack up to the line in the fourth slot and wait for the barcode scanner to align. Wait for the slot light to flash and then slide the rack past the barcode scanner until it clicks into place at the back of the draw. If correctly inserted the light will change to yellow.
30. Gently close the samples draw.

TABLE 47

Elution volumes

Setting up the run:

1. On touchscreen select the samples display, press the ‘batch’ button and then press the ID button. As barcodes are not used for these samples, a unique identifier must be manually assigned to each tube in the samples rack (e.g. 1, 2, 3, 4, etc.). To do this click on the first sample and press the ‘edit ID’ button and then click ‘1’ and then ‘enter’. Repeat this for all samples in the run and then press ‘next’.
2. To assign the correct assay to the samples press ‘select all’ then find the ‘Pathogen’ folder and select the program ‘virus pathogen complex 400 DSP’ from the list. Press ‘next’.
3. To select the elution volume press on the elution rack in slot 1 and then press ‘60’.
4. Press the ‘Queue’ button.
5. Go back to the samples menu and press the carrier button. Select all carrier tubes and then assign the carrier status to them (‘virus pathogen complex 400 DSP’).
6. Press the ‘Run’ button.
7. If prompted to scan any draw again press ‘scan’ and the run will start automatically after completion.

TABLE 48

Break-down of extraction optimisation results