High-throughput Detection of Respiratory Pathogens in Animal Specimens by Nanoscale PCR

No human subjects or experimental animals were used for development of this protocol. Controls were generated by purification of sequence-confirmed amplicons and in vitro transcription for RNA targets. Clinical samples were submitted for routine diagnostic testing to the Cornell Animal Health Diagnostic Center.

1. Plate Design

1. Use primer design software to verify that each assay conforms to standard probe-based real-time PCR cycling conditions with 60 °C annealing using the primer probe test tool (select the quantification probe setting with default parameters).
   NOTE: The representative RNA target used was adapted from the universal influenza A matrix targeted assay published by Shu et al.⁷. The primers and probe are as follows: Forward primer: GACCRATCCTGTCACCTCTGAC, Reverse Primer: AGGGCATTYTGGACAAAKCGTCTA, Probe: Fam-TGCAGTCCTCGCTCACTGGGCACG-QSY. The representative DNA assay used here was adapted from the equine herpesvirus type 1 (EHV-1) detection method published by Elia et al.⁸. The primers and probe are as follows: Forward primer: GCTCTCAGGTTTTACGACATC, Reverse Primer: CTTTACCCAGGCCCTTGAAA, Probe: FAM-TCAACGTGGACAATACCGCAGTGATTAT-QSY.
2. Order nanoscale PCR amplification plates in the desired configuration.
   NOTE: For this study, the 18×3 gene expression format was used (Table 1). Provide sequences for all primers and probes (or inventoried assay IDs) for each target to the manufacturer.

2. Nucleic Acid Extraction

1. Extract total nucleic acid (DNA and RNA) by any desired method.
   NOTE: An automated magnetic bead-based extraction kit was used here according to the manufacturer's instructions (see the materials table for more details).
2. Prepare samples as follows:
   1. Clean the outside of each sample container with 10% bleach and dry thoroughly. Wipe gloved finger tips with 10% bleach between samples to prevent cross-contamination.
   2. Place nasal or deep pharyngeal swabs in any sterile, sealed vial (such as a red top blood collection tube) with several drops of saline added to prevent desiccation prior to processing.
      NOTE: Cotton, plastic, wood-handled, and Dacron and other synthetic swabs are all acceptable, but avoid calcium alginate swabs.
   3. For swabs and tracheal washes, add Dulbecco's Modified Eagle Medium (DMEM) so that there is at least 1-2 ml of liquid. Vortex the swab and DMEM vigorously in the tube. Then, use a pipette to transfer approximately 1 ml of media to a new tube.
   4. Mechanically lyse tissues (100-200 mg in 1 ml of DMEM) using a tissue disruptor according to the manufacturer's instructions followed by centrifugation for 3 min at 825 x g. Transfer 500 µl of the supernatant to a new tube.
   5. For feces, combine 400 mg of feces with 800 µl of 1x phosphate-buffered saline pH 7.4 (PBS). Vortex the suspension for 1 min or longer until the sample is homogenized or completely suspended. Once homogenized, centrifuge the suspension for 10 min at 18,000 x g, then transfer 400 µl to a new tube.
   6. For whole uncoagulated blood, vortex the sample and make a 250 µl aliquot. Add six drops of lytic reagent and vortex to mix. Incubate 15 min at 37 °C.
3. Prepare lysis and wash buffers according to the manufacturer's instructions. Add MS2 phage or an internal control of choice to the lysis buffer as an internal positive extraction control⁹.
4. Add 235 µl of lysis buffer and 175 µl of sample (or PBS for negative control) to each bead tube. Proceed with the manufacturer's protocol.
   NOTE: Purified total nucleic acid was eluted here in 90 µl.

3. Reverse-transcription/Pre-amplification (preamp)

NOTE: Keep all reagents and samples used in the preamp reaction on ice at all times. Following preamp, keep all reagents at room temperature.

1. Assemble eluted DNA and/or RNA, PCR Reaction Mix, nuclease-free water as a negative control, and pooled standards for positive amplification control.
   NOTE: Up to 48 samples can be run on an amplification plate including controls (include at least one negative extraction control for each extraction plate from which samples are to be pulled).
2. Print out a sample map. Have a second person check that the map and sample layout match.
3. In a 1.5 ml tube, add the following to prepare the preamp master mix.
   NOTE: A final volume of 14 µl was used here.
   1. Add a pool of premixed primers from each target such that the final concentration of each primer is 900 nM.
      NOTE: The pool used here was prepared at a 10x concentration, and 1.4 µl added per reaction.
   2. Add random primers to a final concentration of 600 nM or 0.1x.
   3. Add 2x RT-qPCR mix to half of the final volume (7 µl here).
   4. Mix the master mix components together by gently vortexing and spinning down.
4. Perform the preamp in a standard 96-well plate using the left side of the plate only (columns 1-6).
5. Add 8.5 µl of preamp master mix and 5.5 µl of DNA/RNA sample to each well.
6. After combining all reagents (samples, positive controls and negative controls with Preamp mix), thoroughly seal the standard 96-well plate with a clear adhesive seal. Remove any excess seal using a razor blade to prevent formation of seal gaps during cycling. Centrifuge the sealed plate for 20 sec using a PCR plate spinner. Check to ensure all reagents are combined in the bottom of the plate wells.
7. Run the preamp assay on a conventional thermocycler under the following conditions: 15 min at 50 °C; 1 min at 95 °C; 20 cycles of 15 sec at 95 °C, then 2 min at 60 °C; 99.9 °C for 10 min; hold at 4 °C. Program these conditions prior to starting.
8. Turn on the conventional thermocycler, select the preamp cycling program and start the program. Place the 96-well plate in the thermocycler only when the bottom portion of the thermocycler (not the cover) reaches the temperature required for cDNA production (50 °C) by monitoring the temperature reading on the screen. Prior to this, keep the plate cold to minimize the risk of producing false positive results.
9. Once the preamp run is complete, verify that the plate has remained sealed.
10. Proceed immediately to step 4.1. To store this plate overnight, perform the dilution as described in steps 4.2 to 4.5, except instead of loosely covering the plate, seal the plate with a clear adhesive seal and place inside a zipper-lock bag, stored at 4 °C.

4. Dilution of Preamp Plate

1. Remove the appropriate number of amplification plates from the freezer (up to 4 may be run at once). Do not open the packaging. Allow the plate to warm up for at least 15 min at room temperature. Record the serial number and lot number of the plate.
   NOTE: Unopened plates can remain at room temperature for up to 24 hr, but for best practice, remove these from the freezer no more than 30 min before needed.
2. Centrifuge the completed preamp plate for 20 sec using the PCR plate spinner. Turn on the amplification machine and computer and start the associated program.
3. Remove all unnecessary items from a PCR setup area. Put on double gloves and sleeve covers.
4. Remove the seal from the preamp plate and carefully remove and discard the outer gloves. Dilute the preamp products at 1:5 by adding 56 µl of TE buffer to each well of columns 1-6 of the 96-well preamp plate and mixing by pipetting up and down. Go only to the first stop of the pipette, aspirating from the bottom and dispensing higher up but not creating bubbles. Add TE buffer to all 6 columns regardless of unused wells.
5. Reseal the plate with either a clear adhesive or foil seal, and centrifuge it for 20 sec using the PCR plate spinner. Set this plate aside (loosely covered) until step 6.1 is complete. Discard remaining gloves and sleeve covers.

5. Preparation for 384-well Transfer to the Amplification Plate

1. Set up the materials needed to cover and seal the amplification plate: lid, plug, syringe of immersion fluid, plate press, ethanol squirt bottle, and lint-free laboratory wipes. Remove the syringe cap and lock a small plastic tip onto the syringe (requires force). Eject a small amount of immersion fluid onto a paper towel to remove air buildup (it is ok if some small air bubbles remain in the tip). Use the immersion fluid within 60 min of removing the syringe from the vacuum sealed aluminum bag. Once a syringe is opened, do not reattach the cap for later use.
2. Check that the waste bin of the plate loading liquid handler is empty and open the box of tips. Write the expiration date of the tips on the tip box cover when a new box is opened, and ensure that the tips are not expired. Turn on the liquid handler and computer and open the liquid handler software, which will perform a self-test.
3. Click "Setup and Load". Click "Browse" to select the csv file created from the Sample Tracker software. If the file is not shown, go to "Instrument/Edit Preferences" and then click "Change" by "Sample Plate File Folder". Select the folder where the csv file is saved. Then, click "Setup and Load", then "Browse" and select the file.
4. Next, click "Browse" next to the "Plate Holder Position 1" and select the TPF file (provided by the manufacturer) that has the same serial number as the plate.

6. Liquid Handler Transfer

Note: Do not start filling the 384-well plate until all of the above steps are complete. Evaporation is a concern with small volumes — do not let the 384-well plate sit uncovered with liquid inside.

1. Once all of the above steps are completed put on new, well-fitted double gloves and sleeve covers. Then, add 2.5 µl of amplification master mix to a 384-well plate.
2. Transfer 2.5 µl of diluted preamp product to the 384-well plate according to the map in Table 2 and immediately cover the plate tightly with a foil seal. Discard outer gloves and sleeve covers.
3. Centrifuge the 384-well plate for 20 sec in the PCR plate spinner.
4. Do not remove the foil seal, but place the 384-well plate in the liquid handler.
5. Open the package containing an encased amplification plate and place it carefully in the liquid handler in the first slot with the serial number on the right — hold the case by the edges without touching the surface of the plate. Use unwrapped plates within one hour of opening.
   Note: The purpose of the case is to protect the plate from being touched, since this could disturb the small amounts of primers and probes inside the through-holes.
6. Remove the foil seal from the 384-well plate inside the liquid handler once everything is in place. Click Next and then check all the boxes as each step is completed. Click OK to start the run.
7. Close the door to the liquid handler and immediately start the filling process. Stay next to the liquid handler and immediately proceed to the next step once the plate is filled.
8. While the liquid handler is running, remove the protective film from the bottom of a case lid.
   NOTE: The adhesive at the bottom of the case lid is covered by a red tape and a protective film. The film needs to be removed first to access the red tape.
   1. Leave the top film in place until step 6.15. Prime the red tape by pulling slightly to release it from the adhesive.
9. Remove the loaded (filled) amplification plate from the liquid handler and gently place it in the plate press with the serial number on the right.
10. Place the lid on top of the plate with the notched end on the right and adhesive on the bottom (pointing towards the plate).
11. Pull down on the plate press lever carefully closing it; the light will flash for 20 sec. Do not touch the plate press during this time. Once the light turns solid green, lift up the lever carefully and carefully remove the sealed plate by holding it on the edges.
12. While holding the sealed amplification plate vertically by the edges of the case, immediately insert the syringe tip into the loading port at the end of the case, then dispense the immersion fluid slowly in one gentle continuous motion to fill the space between the plate and the lid. Leave one small air bubble in the corner once the plate is immersed.
13. While continuing to hold the plate vertically by the edges, seal the loading port by inserting the plug into the port and twisting the plug clockwise, applying sufficient pressure until the handle breaks off. Grip the edges of the case firmly so that the force of the handle breaking does not cause the case to drop. If a plate is dropped, discard it.
14. Clean the case with a lint-free wipe that has been thoroughly sprayed with ethanol. To dry the case, wipe the case downward with a clean wipe. Gently handle the case; be sure to not apply pressure on the glass above the wells.
15. Bring the sealed plate to the amplification machine. Under the "Instrument" tab, select "Instrument Console." Select the amplification machine then click on the "Open Door" button.
16. Place the amplification plate in the first slot of the plate adapter, checking that the adapter is correctly lined up with A1 in the upper left corner. Orient the plate with the barcode facing up and toward the front of the instrument. Click the "Close Door" button. Note the use of the adapter tray — there is a limit of 10 uses.
17. Under the Home tab at the bottom of the screen, under the Run menu, select OpenArray. Click "Get Plate IDs." The empty boxes will be automatically populated with information about the plate. To begin the run, click "Start Run".
18. Close the liquid handler software and turn off the liquid handler. Place the tip cover over the tip rack. Discard the tips from the waste bin and clean it. Clean and decontaminate all items including the working surface, pipette, waste bucket, and tips boxes.
19. Reseal the preamp plate with a clear adhesive seal and store at -20 °C.

7. Result Analysis

1. Once the run has completed, save the file and then click on the "X" on the tab with the run name at the bottom of the screen to close the file.
2. Click on "Open Door" at the top of the screen to open the door. Remove the amplification plate, checking that no immersion fluid has leaked out. Click on "Close Door" at the top of the screen to close the door.
3. Click "Open" and select the run file to open it. Press the green Analyze button at the top right of the screen. Click "Export" on the left side of the screen and then "Start Export" to export results (as a .txt file). Minimize the file after it opens. Then, click "Export QC Images."
4. Review QC images in the image analysis program according to the following criteria:
   1. Assess loading quality using PRE-READ_CHANNEL_4.tiff and POST-READ_CHANNEL_4.tiff, by checking that there are no dark spots or smudges.
      NOTE: A dye detected by this channel is added by the manufacturer to the primers and probes, which is released into solution when the reaction is loaded. The reading in this channel indicates that reactions were properly loaded and that the primers and probe mixed with dye were present.
   2. Check for leaks or agitation to the plate after loading using S02_C001_t03_p001_m1_x2_e1_cp#_spotfind.tiff. If the image looks dark, increase the brightness (press Ctrl+Shift+C to open controls) until the entire plate is visible. Check that the image looks uniform with no shadows, bubbles, or displaced samples.
   3. Assess lid placement and debris under the lid (bright spots) using STAGE2_CYCLE1_CHANNEL_1.tiff and STAGE2_CYCLE40_CHANNEL_1.tiff.
5. Optional: Open a spreadsheet containing the Results Macro (Supplemental code file). In the exported data file, right click on the tab at the bottom of the screen and select "Move or Copy". In the "to book" field, select "Results Macro.xlsm". Click "Move to end" and check the box "Create copy". Then, Click Ok.
   1. If the Results Macro option does not appear in the "to book" field, simply copy the contents of the exported data file worksheet (click on the upper left cell to highlight all cells and Ctrl-C) and paste it into a new tab.
6. Delete the old "Export" tab and then rename the newly added tab as "Export".
7. Click "Alt-F8" (or click View then Macros) then select "Macro" and click "Run". Then repeat this for Macro2 by clicking "Alt-F8" then selecting "Macro2" and clicking "Run".
8. Rename the Results Macro file using the relevant information (date and serial number), put this information above the table, and save as the same file name in the Exported Results folder. Finally, print out the macro-generated results table.
9. In the amplification machine software, check the curves for each sample and control against the macro table by selecting Analysis/Amplification Plot on the left side of the screen. Then, select the Sample tab, and click on each sample to confirm that there are 2 or 3 positive amplification curves for each of the positive results in the table.
10. Turn off the amplification machine.