A Standardized Procedure for Monitoring Harmful Algal Blooms in Chile by Metabarcoding Analysis

This protocol presents insight into algal blooms and how to monitor toxic algal species effectively. The procedure helps provide an early warning of algal blooms to protect the community from associated damages. Metabarcoding analysis can detect many species in a sample at once, but it requires complicated procedures.

This protocol visually explains each step to help minimize human errors during the analysis. The microscopy procedures will be demonstrated by Henry Cameron from Antofagasta University, sample collection by Jonathan Vilugron from Instituto de Fomento Pesquero, sample pre-treatment by Karen Vergara from Los Lagos University, and the metabarcoding analysis by Ignacio Rilling and Marco Campos from La Frontera University. Begin by collecting three liters of water sample from a target spot.

Filter one liter of the water for 16S rRNA analysis through a tandem filtration to separate the free living and attached bacteria. Next, filter another one liter of water for 18S rRNA analysis through a single filtration with a 0.2 micron membrane. Then, using sterile surgical scissors, cut the filtered membrane in half, wrap one half with aluminum foil for storage, and with the other half proceed to DNA extraction with the Chelex method.

For microscopic analysis, transfer one milliliter of the water sample onto a one milliliter grid slide using a pipette. Observe the sample under a microscope and record the names and quantity of the phytoplankton species. Clean pipettes in the laminar hood cabinet with 70%ethanol, followed by UV exposure for 30 minutes.

For the first PCR amplicon generation, aliquot 22.5 microliters of the master mix in an eight tube strip followed by 2.5 microliters of the DNA sample, then run the first PCR cycle. Next, prepare 100 milliliters of a 2%agarose TBE gel containing 10 microliters of a nucleic acid gel stain. Once the gel is ready, load a mixture containing 1.5 microliters of a DNA loading dye and four microliters of the PCR product and perform electrophoresis at 100 volts for 30 minutes.

Then, image the gel under UV light and confirm the presence of a 500 to 600 base pair band. Missing amplicons can sometimes be solved by diluting the sample to 1, 000-fold with water. Next, use a magnetic beads-based DNA cleanup system to clean the first PCR products, then transfer 20 microliters of each cleaned first PCR product to a new 96-well plate and seal the plate with a Microseal film.

Store the plate at minus 20 degrees Celsius until the next step. For indexation by second PCR, after diluting all index one and index two primers to a one micromolar concentration with PCR grade water in eight tube PCR strips, position the index one primers in a horizontal row and index two primers in a vertical row. Then, using a multi-channel pipette at 2.5 microliters of index one primer to each well of a new 96-well plate.

Next, add 12.5 microliters of a hot start ready formulation to each well. Then, add 2.5 microliters of index two primer and 7.5 microliters of the purified first PCR product to each well. After mixing by pipetting up and down 10 times, cover the plate with a Microseal film and run the second PCR cycle.

After the PCR is complete, place the DNA screen tape in a fragment analyzer. Mix two microliters of sample buffer in three microliters of the second PCR amplicon in new eight tube strips and insert the strips into the fragment analyzer to start the analysis. Ensure that the second PCR amplicons are approximately 613 base pairs for both 16S and 18S rRNA genes.

Purify the second PCR products using a magnetic beads DNA cleanup system, as demonstrated previously. Then measure their DNA concentration using a nucleic acid quantification spectrophotometer. In a new 200 microliter 96-well plate, dilute each purified second PCR product with sterile PCR water to four nanomoles.

Next step onward, the protocol must be performed without halting. Aliquot three microliters of each diluted second PCR product in a new sterile 1.5 milliliter tube to create a pooled library. Always keep the tube at four degrees Celsius.

A day before sequencing, remove a prefilled ready-to-use reagent cartridge from minus 20 degrees Celsius and place it at four degrees Celsius for thawing. On the day of sequencing, set a heat block suitable for 1.5 milliliter centrifuge tubes to 96 degrees Celsius and place the hybridization buffer on ice. Then, dilute molecular grade sodium hydroxide with water and dilute a ready-to-use control library with TE buffer.

Mix 16 microliters of the four nanomole or pooled sample library with four microliters of the control library in a new tube labeled as one. Then, mix 10 microliters of the sample from tube one with 10 microliters of sodium hydroxide in a new tube labeled as two. After vortexing and a brief spin, incubate tube two at room temperature for five minutes, then add 980 microliters of hybridization buffer.

Next, in a new tube labeled as three, mix 260 microliters of the sample from tube two with 390 microliters of hybridization buffer. After mixing the contents by inverting the tube, incubate it at 96 degrees Celsius for two minutes, followed by incubation on ice for two minutes. Next, remove the cartridge from the refrigerator and set up the sample sheet for sequencing with each corresponding index one and index two adapters.

After removing the flow cell from the MiSeq V3 kit, gently clean the flow cell with sterile molecular grade PCR water. Load the entire volume of tube three into the cartridge. Then, in the instrument operation software, select sequencing and follow the instructions to insert flow cell, incorporation buffer, and cartridge.

Finally, load the sample sheet and press Run to start the reaction. This sequence run takes three to five days. 18S rRNA metabarcoding analysis to identify algal species present in sea water collected from Metri Puerto Montt, Chile on February 19, 2019, yielded 13, 750 reads, with over 30 algal species.

The Navicula species was the dominant algo with a relative abundance of 70.73%Also, sufficient abundance was observed for Micromonas, Chaetoceros, Scrippsiella, and Prorocentrum species. The Pseudochattonella species, one of the major causatives of Chilean harmful algal blooms, has a 0.52%abundance. Consistent with the 18S rRNA gene analysis, microscopic observation showed Navicula as the dominant and Prorocentrum as a minor species.

The 12.6%of small, unidentifiable phytoplankton cells recorded by microscopy could be Micromonas, according to the 18S rRNA results. 16S rRNA metabarcoding analysis to identify bacterial species in the same water sample showed 31, 758 reads with over 30 bacterial species. The dominant free living bacterial species was Amylibacter, with a relative abundance of 20.02%followed by Cladella at 13.53%and Algiphilus at 7.06%The relative abundance of two toxic algal species, Alexandrium and Pseudochattonella, is plotted in time cause to find a unique growth pattern.

The relative abundance of all bacterial genus in orders detected from the seawater over five months is plotted and time caused to analyze the population change of certain organisms. It is crucial to be extra clean. Sample contamination should be avoided using an alcohol wipe, UV light, or autoclaving the material.

Diluted solutions should not be reused and tubes should always be capped. Algal detection by metabarcoding must be paired with microscopy. The dominant species can be confirmed by microscopy ensuring no human error during metabarcoding processes.

Harmful algal blooms damage the marine ecosystem and coastal economy. In Chile, algal blooms kill farm salmons, damaging aguacultures. This metabarcoding analysis will increase the efficiency of bloom warnings in Chile.