Low Read Experiment II

This experiment is being repeated due to some errors that occurred in processing the initial plates.

Questions:

1. How can we best accommodate low concentration input DNAs?

   1. Concentrate input DNAs?

   2. Concentrate/normalize low yield products?

   3. Both?

   4. Just put more volume of original DNA in?

2. In the sequence yield, does input concentration, substrate, or some other factor correlate well with low read counts?

Investigations:

Bench(Question 1):

Experimental Sketch of crossed design:

1. Pull aliquots of some high read count and low read count samples. (New low read samples requested)

2. Quantify all samples not including blanks or mock community.

3. For previously low yielding samples, repeat at typical template amount, and at 2x, and 4x volume added to the reaction (in column 2,3,4, and 6, 7, 8). 6x8 samples in low yield, 8 in high yield, 8 of mock community, 8 of blank (ISD only). 72 reactions with different templates. See template below for clarification.

4. Template layout is replicated/duplicated, with different barcodes for duplicates.

5. 1 set of 36 products will not be adjusted for pooling.

6. 1 set of 36 will be adjusted for pooling per qPCR results.

7. Use ISD at lower than typical concentrations for all for PCRs, as a positive internal control and recognizing that this will wreck reconstruction of absolute counts. Will add ISD to master mix.

8. Treat templates with previously high read counts in standard manner, except for lower concentration ISD.

Start Reaction Plate Setup:

• Add Ultrapure H2O to the reaction plates in the following pattern:

NOTE: Water in the master mix has been adjusted to a lower amount. The plate set up below will replace the water usually added to the master mix while also observing if a decrease in water and an increase in template will increase PCR yields.

MasterMix

• Add 5 ul to each well of a hard shell, full skirt plate.

• Add Templates following the pattern below:

• Add 2 ul 1-step primers:

• Seal with bubble seals. Vortex briefly. Spin down.

• Run on Thermocycler Program GSAF36:

MagBead Cleanup:

Some plates were cleaned up using the Nimbus platform protocol “AxyPrep MagBead PCR1 No MM”

Manually, it was done:

• Equilibrate Beads to room Temperature

• Add 24 ul of MagBeads to each well and 15 ul of replicate to same well of replicate

• Pipette mix up and down 10 times.

• Incubate at RT for 5 minutes

• Secure plate on magnet plate; incubate at RT for 5 minutes (until wells are clear)

• Remove 65 ul from each well; keep tips to left or right depending on the column to avoid bead pellet.

• Add 100 ul Fresh 80% EtOH to each well. Incubate 30 seconds. Remove 100 ul from each well

• Add 100 ul Fresh 80% EtOH to each well. Incubate 30 seconds. Remove 100 ul from each well

• Reaspirate from each well to assure maximum EtOH removal

• Allow plate to air dry for 7 minutes.

• Remove sample plate from magnet plate.

• Add 40 ul TE; pipette mix 10+ times. Incubate 2 minutes at RT.

• Place sample plate back on magnet for 5 minutes or until all wells are cleared.

• Transfer 40 ul to labeled transparent plate (Plate name_PCR_MIDs)

qPCR all products:

• Make 1:1000 dilutions of all samples from the PCR plates by adding 1 ul to 999 ul TE in a deep well plate:

• Add 16 ul of Illumina Library Quantification MasterMix to each well:

• Add 4 ul of template, pool, or standards to each well:

Results:

Pool all samples:

Blue samples (top half of plate) will be pooled at 1* ul per sample.

*The original experimental design was to add 2uL per sample for the blue pool but with the extremely low yields occurring in the red half of the plate I reduced the pooling amount of the blue pool to make the two pools closer in total concentration.

Red samples (bottom half) will either just be pooled per qPCR numbers or concentrated via SpeedVac, reconstituted to a higher concentration, and pooled by qPCR results.

NOTE: Row H showed significantly lower yields. I believe this could have occurred due to evaporation of the PCR master mix. The master mix was prepared ahead of time, plated, and refrigerated until samples could be loaded. We prepare master mix plates ahead of time often, with no noticeable evaporation, but I did notice a slight decrease in volume on the outer edges of the plate during sample loading. Due to the master mix being specialized for this experiment, I proceeded with the plates as is since they were in duplicate. If the following steps below do not show the desired result, we could repeat the experiment with the master mix preparation and sample loading occurring on the same day.

Calculations for Red Pool:

Since the majority of the samples in the red pool were in the 10-99 nanomole range, the following samples will be added in higher quantities to ensure the samples are pooled in the same order of magnitude:

In theory, if you decrease the elution volume by 50%, the DNA concentration should increase by 50%. That being said, in order for some of the samples to reach the desired concentration, we will need to vacuum concentrate and resuspend at a lower elution volume.

• samples with concentrations between 10-99 will have 1uL added to the pool.

• samples highlighted in red were too low to normalize to 10 nanomoles so I just resuspended them at 10uL and used half to at least increase the input quantity.

qPCR blue/red pools:

• Make 1:1000 dilutions of blue and red pools by adding 1 ul to 999 ul TE in a 1.5mL tubes.

• Run each pool in triplicate.

• Add 16 ul of Illumina Library Quantification MasterMix to each well:

• Add 4 ul of template, pool, or standards to each well:

Results:

Average Results:

LR_Blue_Pool: 18.10 nanomoles

LR_Red_Pool: 8.36 nanomoles

iSeq Run

Dilute to 1 nM based off qPCR results. qPCR results are in pM, but 1:1000 dilution used. The results are effectively in nM for pool.

*Due to the blue pool only having a total of 36uL, the 1nM pool calculations had to be at a total volume of 500uL instead of 1000uL.

Blue Pool

• 500/Results = ul of Pool to Add

500/18.1= 27.5 uL of Pool to Add

• 500- ul of Pool to Add = ul of “10 mM Tris 8.5” to Add

500- 27.5 = 472.5 uL of 10mM Tris 8.5

Red Pool

• 500/Results = ul of Pool to Add

500/8.36 = 59.8 uL of Pool to Add

• 500- ul of Pool to Add = ul of “10 mM Tris 8.5” to Add

500- 59.8= 440.2 uL of 10mM Tris 8.5

Combine Blue and Red pools in equal parts once pools are at 1nM

Dilute 1 nM full pool to loading concentration of 50 pM:

• Add 5 ul 1 nM Pool to 85 ul “10 mM Tris 8.5” and 10 ul 50 pM PhiX

• Remove iSeq 100 i1 Flow Cell from refrigerator 5’s crisper drawer and open white foil pack and allow to equilibrate to RT for 10-15 minutes.

• Open “iSeq 100 i1 Reagent Cartridge v2”. Turn on iSeq100

• Click on “Sequence”. Watch Video. Do what video tells you to do. Follow on screen instructions until run starts.

• Results located: Data/SequencingRuns/”folder with applicable date”/Alignment_1/Fastq/*.fastq.gz

Data wrangling(2):

Use R to associate NS2 input concentrations with sample names, read counts, and % ISD reads