16S rRNA gene Library Preparation Protocol

UV the MPure machine before use.

Clean MSC with DNAZap.

Remove samples to be extracted from the freezer, to thaw in the fridge (4°C).

Once thawed vortex for 0h 0m 10s.

Proceed in MSC.

A negative control (e.g. nuclease free water), should be done for each kit (approx. every 50 samples) to show the presence of contaminants (particularly important in low biomass samples) and records should be kept to link this to the samples extracted, machine used, and kit information.

Pipette mix and remove up to 800µL from the storage tube and transfer to a labelled 1.5 ml Eppendorf tube

Centrifuge tube for 0h 10m 0s at 13200rpm.

Remove and discard all bar 100µLof the supernatant. Resuspend the bacterial pellet in the remaining liquid.

Add 600µLof BLB2 buffer (MPure Bacterial DNA Extraction Kit’, MP Biomedicals)and vortex to further resuspend.

Transfer the volume to a labelled lysis tube ( Lysing Matrix E, MP Biomedicals). Label the side of the tubes.

Place lysis tubes in the ‘FastPrep Instrument’ and run for 0h 0m 40s at speed 6000.

Centrifuge tube for 0h 10m 0sat 13500rpm,0h 0m 0s.

Set Up the MPure 12 Reagents and plasticware:

Place the reagent cartridge in the large flat block at the back, it should click into place, and the writing and release flap should be facing toward you.

The clear plastic mixing wells are clipped in front of these.

Then place the black plastic container set into the next slots, into the first of these place a plastic pin, in the third place a tip.

Put an open labelled elution tube in the closest holes in the metal rack (labelled E).

Remove the supernatant (450µL) from each lysis tube to a labelled blue sample tube from MPure bacterial DNA extraction kit.

Place these in the metal rack, in the slots labelled “S” on the machine. 

Machine instructions: 

Select “START”, it will scan the setup from the sheet in the kit. 

Select the protocol : Mpure Bacterial DNA extraction kit with barcode: “OP02006”·       

Select sample volume 400ul with barcode ”SV0400”·       

Select elution volume 50ul with barcode “EV0050”·       

Press Enter to start the extraction 

The run time is around 1 hour. The machine will stall if an error develops, but will not stall if a tip is lost, or plasticware comes unclipped.

Remove elution tubes, checking a reasonable volume is present, and store these at -80°Cin a labelled box.

Clean MSC or PCR hood with DNAZap.

Dilute primers to working concentration (10millimolar (mM),) in NFW, if there are no pre-existing dilutions < 2 weeks old.

Aliquot diluted primers into labelled, linked PCR tubes. Freeze upright overnight if necessary.

Rack out the samples for the library into 12 x 8 rack according to use, complete plate plan, put on lid, label and store in -80°C.

Remember to include:

• PCR negative control(s) from the relevant DNA extractions

• positive control (mock community or bacterrial DNA)

• negative PCR control (nuclease free water).

• any diluted, or pre-spiked samples.

Wipe down MSC with Distel/DNAZap, and then 80% ethanol.

Wipe down all equipment with distel and put in the MSC. 

Close MSC, and UV.

Defrost the reagents on ice.

Add 14.25µL nuclease free water to labelled master mix tubes.

Add 5µLQ5 buffer (Q5 High-Fidelity Polymerase Kit from New England Biolabs) to the master mix tubes. Mix before use, vortexing if salts have fallen from solution.

Add 10millimolar (mM) dNTP (0.5µLper well) to the master mix tubes. Mix before use.

Pipette mix the master mix, and keep on ice.

Record reagent lot numbers on the library plate plan sheet. Try to use the same PCR reagents for all samples, if possible.

Add Q5 enzyme (5µLper well) to individual master mix tubes immediately before use, keep on ice. Pipette mix.

Prepare a 96 well plate for use, labelling and adding orientation points as necessary. Use the plate ice block from this point on.

Pipette 20 μl of Master mix into each well of the 96 well plate. There is no need to change tips, but the buffer may bubble.

1.25 μl of each primer (501-508) can be transferred using a multichannel pipette in the vertical from the set of linked PCR tubes, swap tips if the plate or tubes are touched. Check each tip has taken up the correct volume.

1.25 μl of each primer (701-712) can be transferred using a multichannel pipette on the horizontal from a set of linked PCR tubes, tips should be changed after each use. Check each tip has taken up the correct volume.

Apply a plate seal and move the plate, and it’s chiller unit into a second MSC if available.

Retrieve the samples from the fridge, spray the closed rack into the MSC.

Carefully remove the plate seal. Change gloves.

Add 2.5 μl of each sample DNA template into each well, according to your plate plan. Change tips every time and discard if they have touched anything before use.

Return samples to the fridge.

Apply a plate seal and use a roller and squeegee to ensure that the wells seal properly. If you find large amounts of loss from outer wells apply autoclave tape around the edges.

Run this plate on a PCR machine with a heated lid.

Initially at 98°C for 2minutes.

32 cycles of 98°C for 30 seconds, 50°C for 30 seconds, 72°C for 1 minute and 30

seconds.

Finally 72°C for 5mins and hold at 4°C.

Wipe Down MSC and pipettes with DNAZap.

Repeat steps 28-40 until the library has been prepared and run in triplicate.

These plates can be frozen at -20°C before clean up.

Bring beads to Room temperature, prepare 80% ethanol and TRIS-HCl before beginning.

Defrost plates on ice and spin down before opening. Change gloves.

Combine the triplicate PCR products into 1 plate using a multichannel pipette.

Vortex the beads stock (Agencourt AMPure XP beads) for 30 seconds to ensure even dispersal. Pour into a trough.

Add an equal volume of beads per volume of PCR product (probably 75µL) to each well, gently pipetting up and down x10, changing tips for every column.

Incubate at Room temperature for 0h 5m 0s

Move the plate to the magnetic stand for 0h 2m 0suntil the supernatant has cleared.

Leaving the plate on the magnetic stand use a multichannel pipette to remove and discard the supernatant, changing tips every time.

If you disrupt the beads leave the well until the end to allow them to resettle. The beads should remain attached to the side of the well.

Leaving the plate on the magnetic stand wash the beads with 200µLfresh 80% ethanol, using a multichannel pipette.

Incubate on the magnetic plate for 0h 0m 30s. Carefully remove and discard the supernatant into a waste container.

Leaving the plate on the magnetic stand wash the beads with fresh 80% ethanol. Using a multichannel pipette add 200µL of fresh 80% ethanol to each well.

Incubate on the magnetic plate for 0h 0m 30s.Carefully remove and discard the supernatant into a waste container. Use a P20 with fine tips to remove any residual ethanol.

Leave the PCR plate on the magnetic plate for 0h 10m 0s to air dry.

Remove the plate from the magnetic stand. Using a multichannel pipette add 27.5µLof TRIS-HCl pH8 into each well of the plate.

Pipette up and down x10 to re-suspend the beads, Some wells may need more flushing than others, ensure that the beads are fully resuspended.

0h 2m 0s Incubate the plate at room temperature for 0h 2m 0s

Put the PCR plate on the magnetic stand for 0h 2m 0s (or until the supernatant clears).

Using a multichannel pipette move 25µL from each well into a new clean labelled PCR plate. Change tips between samples. Do not disturb the bead pellet .

Seal plate thoroughly. If freezing at -20°C use aluminium foil. Keep at 4°C overnight.

Label Qubit tubes with the well names (Sample Tubes).

Label two Qubit tubes 'S1' and 'S2' and place 10µLof Qubit HS Standard 1 into the first and 10μl of Standard 2 into the second RTP.

Briefly vortex and spin down the samples.

Wrap tape and foil around a 50ml Falcon Tube to ensure no light enters.

Into the Falcon Tube, add Qubit HS Buffer (199µL x (sample no.+2)) and Qubit HS Reagent (1µL x (sample no.+2)), close and vortex well. This working solution is light reactive; ensure that the duration of light exposure is minimal.

Add 190µL of Working Solution to each of the Standard Tubes, vortex for 0h 0m 5s and leave for 0h 2m 0s. Ensure there are no bubbles and all liquid is at the bottom of the tube.

Select dsDNA HS on the Qubit machine & calibrate using the 2 Standard Tubes. Note down the values for the Standards.

Vortex the Working Solution Tube and add 198µLnto a Sample Tube, then add 2µL of the correct sample, vortex for 0h 0m 5s and leave for 0h 2m 0s. Ensure there are no bubbles in the Sample Tube and all liquid is at the bottom of the tube.

Place Sample Tube into the Qubit machine and record the DNA concentration.

Repeat steps 68-69 for the remaining samples. Dilute any samples where the concentration is too high and repeat to get a value.

Find the highest and lowest concentration of pooled PCR product & use this to decide the amount of DNA from each sample to be added to equimolar mix (ensure volume of lowest is possible (eg. 20µL) and volume of largest is possible 1µL). Remember to scale up any diluted samples.

e.g.             Highest = 55.0μg/ml         Lowest = 6.2μg/ml

55μg/ml  x 3 ul = 165μg              165 / 6.2 = 26.6μl

However, only 20μl any sample, so: 55 x 2 = 110μg              110 / 6.2 = 17.7μl)

Once amount of DNA is calculated, use the following formula to work out volume of each sample to be added to equimolar mix: Volume (μl) = DNA amount (μg) / Sample Concentration (μg/ml). If the concentration of a sample is <2ng/μl, add all the sample volume to the equimolar mix to give the best chance of enough reads.

Print list of needed volumes in 96 well format.

Mix the correct volumes of each pooled PCR product to create the equimolar mix in a labelled, screw cap 2ml eppendorf. Add the largest volumes first.

Pipette mix and check that the total volume is correct.

Split the equimolar mix volume in half.

Store half of the equimolar mix at -20°C in a o-ring screwcap tube.  

Mix the unfrozen half of the equimolar mix with loading dye as instructed by the dye information leaflet.

Make 1% agarose, 1x TBE gel with the nucleic acid gel stain of your choice, load into buffer tank and top up the buffer until it is level with the top of the gel.

Load the Equimolar mix into the gel, filling the well(s) as much as possible.

Run for 0h 10m 0sat 90V with the buffer level with the top of the gel. The equimolar mix should have moved into the gel.

Add buffer to cover and run for approximately 1h 30m 0s.

Use a UV transilluminator to visualize the bands. Cut out the bands with as little excess gel as possible. Wear suitable face and eye covering, and ensure any nearby workers are taking suitable precautions. Dispose of any sharps appropriately.

Weigh as many 1.5ml tubes as you have bands, and note their weight. Place the bands in to the 1.5ml tubes. Weigh the tubes to estimate the weight of gel slice.

Add 10µL of Membrane Binding Solution (Wizard SV Gel and PCR Clean Up Kit from Promega) per 10mgof gel slice and vortex briefly.

Eg. tube + gel slice = 1.48g, tube = 1g, gel slice = 0.48g, add 480μl of Membrane Binding Solution.   

Place tube into a heat block at 55°C until gel dissolves completely. You may want to vortex during this incubation to hasten the dissolving.

There will only be 1 collection tube used per Equimolar mix. Place a Minicolumn into a Collection Tube and pipette up to 700µLof dissolved gel into the column.  Incubate for 1 minute.

Centrifuge the SV Minicolumn at 13200rpm,0h 0m 0s for 0h 1m 0s.

Discard the supernatant into a labelled liquid waste container and reinsert the SV Minicolumn into the Collection Tube.

Repeat steps 87-89 until no dissolved gel mixture remains ( if > than 700µLof dissolved gel).

Add 700µL of Membrane Wash Solution (add ethanol before use as instructed by kit).

Centrifuge at 13200rpm,0h 0m 0s for 0h 1m 0s.

Discard the supernatant and reinsert the SV Minicolumn into the Collection Tube.

Add 500µL of Membrane Wash Solution.

Centrifuge at 13200rpm,0h 0m 0s for 0h 5m 0s

Discard the supernatant. Centrifuge the tube and minicolumn at 13200rpm,0h 0m 0s for 1 minute.

Transfer the SV Minicolumn to a 1.5ml tube and add 100µLof Nuclease-Free Water directly onto the membrane, incubate for 0h 1m 0sat RTP.

Centrifuge at 13200rpm,0h 0m 0s for 0h 1m 0s.

Check the water has gone through the column and that the membrane is dry. If water remains on the matrix, repeat step 97.

Discard SV Minicolumn and split sample, for storage in a labelled screwcap tube with O-ring. Keep half as a spare and submit the other half for sequencing.