Efficient Nucleic Acid Extraction and 16S rRNA Gene Sequencing for Bacterial Community Characterization

The overall goal of this procedure is to isolate high-quality DNA and RNA from swabs and then determine their bacterial composition using 16S rRNA gene sequencing. This method can help a researcher identify which bacteria are contained within a given human sample and determine associations between particular bacteria and host characteristics or disease status. Though this method was designed to provide insight into human vaginal bacterial communities the method can also be applied to study bacterial communities from other body sites, such as the nares, mouth, skin, and rectum and from other organisms.

Note that this video highlights the trickier parts of the protocol and does not cover the sequencing portion or the sequencing analysis. Brittany Bowman, a technician from my laboratory, will demonstrate. Begin this procedure with ectocervical swabs collected and stored on wet ice as described in the accompanying document.

In preparation for nucleic acid extraction, clean all of the surfaces of the hood and all of the items brought into the hood with bleach followed by a decontaminant that removes RNases, DNases, and DNA. In the hood, place one bead beading tube per sample. To each tube, add 500 microliters of NaCl-Tris-EDTA buffer, 210 microliters of 20%SDS, and 500 microliters of phenol chloroform isoamyl alcohol.

Careful handling of the swabs is critical to obtaining clear results with this protocol. It takes a little practice to transfer the swab without dropping it and it is easier if you use a hollow shafted swab like this one. Using sterile forceps, transfer the swab from the transport vial into the bead beading tube.

Thoroughly rub the swab head against the internal walls of the tube for at least 30 seconds. Place the tube on ice. Then change gloves and repeat the swabbing procedure with the next sample.

Once all of the samples have been rubbed vigorously allow them to chill on ice for at least 10 minutes. After 10 minutes have passed to remove the swab, hold the handle with sterile tweezers and, using a clean P200 tip, press the head against the inside of the tube wall. This process with liberate liquid from the swab, increasing nucleic acid recovery.

Place the bead beading tube into the bead beader and homogenize the sample for two minutes at four degrees Celsius. Then, centrifuge the bead beading tube at 6, 000 gs and four degrees Celsius for three minutes to pellet the debris and separate the aqueous and phenol phases. After the spin, three phases can usually be seen.

The top layer is the aqueous phase, containing the DNA and RNA. The bottom layer is the organic phase, containing lipids and debris from the cells and swabs. The thin white middle layer, called the interphase, contains protein.

Transfer the aqueous phase, about 500 to 600 microliters, to a sterile 1.5-milliliter tube. Add an equal volume of phenol chloroform isoamyl alcohol. Mix by inversion and brief vortexing.

Centrifuge the tube for five minutes. After the spin, transfer the aqueous phase to a new sterile 1.5-milliliter tube. Take care not to transfer material from the interphase or the underlying phenol phase.

Note the volume transferred and save the phenol phase for future protein isolation. Next, add 0.8 volume of isopropanol and 0.1 volume of three molar sodium acetate. Mix thoroughly by inversion and briefly vortexing.

Precipitate the nucleic acid by chilling the tube at 20 degrees Celsius for at least two hours and as long as overnight. Following precipitation, centrifuge the tube for 30 minutes. After the spin, carefully use a pipette to remove the supernatant, leaving the pellet intact.

Add 500 microliters of 100%ethanol. Dislodge the pellet with gentle vortexing or pipetting without touching the pellet. Centrifuge for five minutes.

Using a P10 pipette, remove and discard as much ethanol as possible without disturbing the pellet. Air dry the pellet at room temperature for 15 minutes. Resuspend the pellet in 20 microliters of UltraPure 0.1X-Tris-EDTA buffer.

Allow the sample to chill on ice for 10 minutes and pipette repeatedly to ensure full resuspension. Measure the nucleic acid concentration using a spectrophotometer. If desired, separate DNA from RNA using a column cleanup kit following the manufacturer's protocol.

The PCR setup is a critical step in this procedure. It's very important to minimize contamination by ensuring sterility of the reagents, the PCR hood, and the gloves. Work quickly but carefully and use proper technique when moving items into and out of the hood.

Place a clean benchtop cooler rack for microcentrifuge tubes and a PCR plate cooler in the hood. Label 8-well strip tubes with individual caps and place them in the PCR cooler. Amplification should be performed in triplicate and a no template water control should be included for each primer pair.

Place a microcentrifuge tube in the cooler for the master mix. Then, for each sample, combine UltraPure water, 5X HF buffer, DNTPs, 515F forward primer, and 3%DMSO as described in the accompanying document. Then, add polymerase to the master mix and mix by pipetting.

To the first well, add 90 microliters of the master mix and two microliters of the reverse primer, mix well. Then to the fourth well, which serves as the no template control, add 23 microliters of the master and two microliters of water and mix. Next, add six microliters of the appropriate sample to the first well.

Mix thoroughly and transfer 25 microliters to the second well. Change tips, then transfer another 25 microliters from the first well to the third well. Firmly cap every well.

Take care not to touch the inside of the wells or the cap. Then, repeat this process for each sample. Once all of the samples are prepared, quickly spin the strip tubes, transfer them to a thermocycler, and run the program.

After the run is complete, quickly spin the tubes to collect liquid from the walls. Then, on a clean lab bench, combine the triplicate PCR reactions from each sample in a labeled sterile microcentrifuge tube. Transfer 25 microliters of each no template control into a separate sterile tube.

Do not combine amplicons from different samples yet. To validate successful PCR amplification, load five microliters of each sample on a 1.5%agarose gel and perform electrophoresis at 120 volts for 30 to 60 minutes. View the gel under UV light.

Verify successful amplification of each sample by noting a single strong band around 380 base pairs. If there is a double band, re-amplify the sample with a different reverse barcode. If there is no band at all, re-amplify the sample using either the same reverse barcode or a primer with a new reverse barcode.

If re-amplification is unsuccessful, PCR inhibitors may be present in the sample, in which case, perform a column-based DNA cleanup to remove PCR inhibitors. Store the remaining 70 microliters of amplicon at 20 degrees Celsius. Discard the remaining 20 microliters of the no template control, assuming it did not yield a band.

To create the amplicon pool combine two to five microliters of each amplicon in a single sterile tube. If the band from a sample is weak add twice the volume relative to the rest of the samples. Then remove the PCR primers using a kit.

If applicable, combine the primer-free amplicon pools to create the final library. Determine the DNA concentration of the library using a spectrophotometer or a fluorometric system. Finally, dilute the samples to two nanomolar and send them for sequencing.

Detailed information about sequence analysis is provided in the accompanying document. High-quality DNA and RNA was isolated and the DNA was used as the template for PCR amplification of the 16S rRNA gene V4 region as described in this video. Gel electrophoresis was used to confirm the presence of a single band around 380 base pairs in every sample that was amplified with template.

The no template controls run in parallel with the same primer pair did not have a band present. The PCR amplicons were then combined into a single tube, quantified, and further diluted before sequencing. Shown here is a representative bar plot of the sequence quality scores at each position of the read.

It is normal for the sequence quality to drop after 200 base pairs. But the average quality score should remain above 30. After watching this video, you should have a good understanding of how to isolate DNA and RNA from swabs using a phenol chloroform-based extraction, amplify the V4 region of the 16SR ribosomal RNA gene, and prepare a library for high-throughput sequencing.

Once mastered, this technique can be done in two days if it is performed properly. While attempting this procedure, it's important to take great care to prevent contamination during sample collection, nucleic acid extraction, and PCR amplification. Don't forget that working with phenol chloroform can be extremely hazardous and precautions such as working in a vented hood and wearing impervious gloves, safety glasses with side shields, and a lab coat should always be taken while performing this procedure.