Characterizing Microbiomes via Sequencing of Marker Loci: Techniques To Improve Throughput, Account for Cross-Contamination, and Reduce Cost

ABSTRACT New approaches to characterizing microbiomes via high-throughput sequencing provide impressive gains in efficiency and cost reduction compared to approaches that were standard just a few years ago. However, the speed of method development has been such that staying abreast of the latest technological advances is challenging. Moreover, shifting laboratory protocols to include new methods can be expensive and time consuming. To facilitate adoption of new techniques, we provide a guide and review of recent advances that are relevant for single-locus sequence-based study of microbiomes—from extraction to library preparation—including a primer regarding the use of liquid-handling automation in small-scale academic settings. Additionally, we describe several amendments to published techniques to improve throughput, track contamination, and reduce cost. Notably, we suggest adding synthetic DNA molecules to each sample during nucleic acid extraction, thus providing a method of documenting incidences of cross-contamination. We also describe a dual-indexing scheme for Illumina sequencers that allows multiplexing of many thousands of samples with minimal PhiX input. Collectively, the techniques that we describe demonstrate that laboratory technology need not impose strict limitations on the scale of molecular microbial ecology studies. IMPORTANCE New methods to characterize microbiomes reduce technology-imposed limitations to study design, but many new approaches have not been widely adopted. Here, we present techniques to increase throughput and reduce contamination alongside a thorough review of current best practices.


Introduction
This standard operating procedure (SOP) was inspired by the approach developed by the Genome Sequencing and Analysis Facility at the University of Texas (https://wikis.utexas.edu/display/GSAF/Home+Page).
This SOP is for amplification of the 16s (515-806 primer pair) and ITS (ITS1f-ITS2 primer pair) loci from environmental samples, but it could be modified to amplify many other marker loci, simply by changing primer sequences and possibly making minor modifications to the temperatures used during PCR. Libraries made with this protocol are to be sequenced on Illumina platforms, including the iSeq, MiSeq, HiSeq, and NovaSeq.
The SOP makes use of 96 unique "coligos" for tracking cross-contamination among wells (see main text). We also use a synthetic DNA internal standard (ISD) inspired by (Tourlousse et al. 2017). We suggest that future iterations of this SOP be amended to include an internal standard comprising several unique sequences (see Harrison et al. 2020).
Users should be aware that coligo reads will have poly-G tails that arise from a lack of signal from the sequencing machine. These tails are readily trimmed off during bioinformatics or used as a way to identify and remove coligos from the dataset. We also note that coligos, due to their short length, do not always merge well. Therefore, when performing analyses to determine the extent of cross-contamination we recommend using forward reads only. Alternatively, coligos can be synthesized such that they are longer and will merge more readily, however this will increase the cost of coligo synthesis.
For the two-step procedure, primer design is the same except only a part of the Illumina flow cell adapter is included.

Protocol
We present two protocols here, a one-step procedure that adds Illumina flow cell adapters and barcodes during template amplification and a two-step procedure that allows for the use of shorter oligos but requires two rounds of PCR (for visual description see main text). The two-step procedure adds a portion of the Illumina flow cell adaptor and the barcode during initial priming and amplification of the template. A second round of PCR adds the remaining portion of the Illumina adaptor.
The two-step procedure is commonly used because it requires short, inexpensive oligos. However, it takes longer to complete and, for best results, requires an additional PCR clean-up step, which adds significant cost. We have shifted to a one-step approach which has a much lower per-reaction cost and requires less time. The one-step approach requires longer oligos than the two-step technique and these longer oligos incur a large initial cost of a few thousand USD (at the time of writing). However, for laboratories processing many samples, this large initial cost will rapidly be defrayed due to time and consumable savings (e.g., in magnetic bead clean-up kits, pipette tips, and so on).

A note on cleanliness
Every effort should be made to reduce contamination and keep a clean work space. The work bench should be cleaned with a solution that degrades DNA between projects. Nucleic acid extraction and sample preparation (e.g., the weighing of soil) should be done in a different room from PCR, if at all possible, to minimize the chance of contamination. If a different room is not available, then at least perform extractions and PCR on separate benches and clean all equipment thoroughly when shifting from performing extractions to performing PCR. Pipettes and other equipment should be cleaned daily, or more often, as needed. Ideally, PCR should be performed under a hood. Gloves should be worn during sensitive steps, such as when reagents and MID-plates are opened. Gloves must be clean, one cannot touch anything that has not been cleaned and expect the gloves to perform their function (e.g., cabinet handles, writing tools, keyboards, if they haven't been cleaned then one's gloves will be contaminated). Do not pipette directly from stock solutions.
Follow all other laboratory rules and keep careful notes. Labeling of samples is the most important step of this entire process. Without proper labels all is lost! 3.2 One-step procedure 1. Aliquot 30 µL of full concentration extracted DNA into a new 96 well plate (unskirted, standard well depth). Save this plate in the event the library needs prepared again. Label the plate. We currently advise performing PCR in duplicate.
2. Add 6 µL of control oligo pool to environmental DNA aliquot. The control pool includes 16S and ITS coligos (0.01 pg µL −1 of each) and 0.03 pg µL −1 each of the 16S and ITS internal standards. If desired, quantify DNA concentration for each sample and normalize. Currently, we are normalizing to 10 ng µL −1 . Save aliquot of DNA in case the library needs to be prepared again. Label the plate.
3. Add 11 µL of Master Mix to each well of a new plate (see Table 1 for Master Mix ingredients). Label the plate. 6. Apply the following PCR recipe: 2. Add 6 µL of control oligo pool to environmental DNA aliquot. The control pool includes 16S and ITS coligos (0.01 pg µL −1 of each) and 0.03 pg µL −1 each of the 16S and ITS internal standards. If desired, quantify DNA concentration for each sample and normalize. Currently, we are normalizing to 10 ng µL −1 . Save aliquot of DNA in case the library needs to be prepared again. Label the plate.
3. Add 7 µL of Master Mix #1 to each well of a new plate (see Table 3 for Master Mix ingredients). Label the plate.
5. Add 2 µL of template DNA (normalized) to each well.
6. Apply the PCR recipe shown in Table 4.  8.13 Remove the liquid from the well and place in a new, labeled plate. This is the cleaned DNA.
8.14 Prepare flow cell Master Mix (Table 5). 11. Apply PCR recipe #2 (Table 6) to this plate (which contains the flow cell Master Mix and the cleaned template from PCR #1). 12.5 Secure plate on magnet; incubate at room temperature for 5 minutes (until wells are clear).
12.6 Remove 65 µL from each well, while avoiding the bead pellet, which contains the DNA.
12.7 Add 100 µL fresh 80% ethanol to each well and incubate 30 seconds then remove 100 µL from each well.
12.9 Aspirate from each well again to assure maximum ethanol removal. Ethanol can interfere with PCR.
12.10 Allow plate to air dry for 7 minutes. This allows remaining ethanol to evaporate.
12.11 Remove sample plate from magnet plate.
12.13 Place sample plate back on magnet for 5 minutes.
12.14 Transfer 30 µL to a clean PCR plate.
14. Optional: Normalize all samples to 9 ng µL −1 prior to sequencing. This step is not needed if high sequencing depth is likely since sufficient data should be obtained from all samples.
15. Optional: Check molar concentration of the pooled library via qPCR.
16. If sequencing is to be performed in days or weeks then store library in refrigerator. If sequencing will not occur for awhile then store the library in a non-cycling freezer.