Standard Genetic Methods – Metabarcoding Standard Operating Procedures

Katrina Lohan; Ruth DiMaria; Calli Wise; Tara Sill; Lael Collins; Emma Palmer

Jun 29, 2026

Standard Genetic Methods – Metabarcoding Standard Operating Procedures

DOI

https://dx.doi.org/10.17504/protocols.io.kxygxrp3wg8j/v1

Katrina Lohan¹,
Ruth DiMaria¹,
Calli Wise¹,
Tara Sill¹,
Lael Collins¹,
Emma Palmer¹

¹Smithsonian Environmental Research Center, Coastal Disease Ecology Lab

Coastal Disease Ecology Lab - SERC

Calli Wise

DOI: https://dx.doi.org/10.17504/protocols.io.kxygxrp3wg8j/v1

Protocol Citation: Katrina Lohan, Ruth DiMaria, Calli Wise, Tara Sill, Lael Collins, Emma Palmer 2026. Standard Genetic Methods – Metabarcoding Standard Operating Procedures . protocols.io https://dx.doi.org/10.17504/protocols.io.kxygxrp3wg8j/v1

License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited

Protocol status: Working

Created: June 25, 2026

Last Modified: June 29, 2026

Protocol Integer ID: 319832

Keywords: Metabarcoding, standard genetic method, sanger sequencing standard operating procedure, metabarcoding standard operating procedure

Abstract

Standard Genetic Methods – Sanger Sequencing Standard Operating Procedures 

Guidelines

**New users must be trained by a technician or post-doc! **

Whenever in the lab, all individuals must wear shoes that completely cover their feet.

Whenever working with any chemicals, reagents, or DNA, all individuals must wear clean nitrile gloves and are responsible for understanding any hazards associated with those chemicals (Safety Data Sheet; SDS).

Wear gloves when working with bleach and work in a well-ventilated area.

This protocol was developed with the assumption that the end-user possesses baseline knowledge of appropriate aseptic/clean techniques and a comprehensive understanding of the potential risks of contamination inherent to the workflow.

*If there are any questions or concerns, please do not hesitate to ask Ruth DiMaria or Katrina Lohan*

I. DNA EXTRACTIONS

• WORK IN PRE-PCR DESIGNATED SPACES ONLY
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILTER (BARRIER) PIPETTE TIPS
• CREATE ALIQUOTS OF REAGENTS TO PREVENT CONTAMINATION OF STOCK BOTTLES
• For samples and reagent bottles, only open one tube at a time. Do not touch the inside of the tubes with gloves. If you do so, CHANGE GLOVES immediately.
• Additional steps and modifications must be followed during pre-PCR tasks if the project targets bacteria, viruses, or human pathogens. Contact Ruth about the training required.

*Follow all safety precautions outlined in instruction manuals for the appropriate kit

*Choose kit appropriate for sample type – animal tissue, culture, sediment, water

*Always wipe down areas with diluted bleach (10%) before beginning. First wipe down the surface with tap water before applying diluted bleach as a precaution against a negative chemical reaction with any residual guanidine hydrochloride or other incompatible chemicals commonly used in the lab. For more information, refer to the Sodium Hypochlorite (Bleach) Safety Fact Sheet

*This protocol is extremely sensitive and highly prone to cross-contamination. Thus, it is imperative that tubes and bottles not be left open. 

For reagents, remove the lid, pipette out any liquid needed, and place the lid back on top. The lid does not have to be secured in place until it is no longer needed, but it must be covered when not imminently in use.

For samples, only open one tube at a time. Do not touch the inside of the tubes with gloves. If you do so, CHANGE GLOVES immediately. 

*UV-crosslink all tubes (if not already sterile from packaging) before beginning (2 minutes standard).

*All waste (solid or liquid) containing hazardous materials should be disposed of in an appropriate labeled container, tracked and stored in the Satellite Accumulation Area (SAA). Contact Ruth DiMaria or Calli Wise for required training. Guanidine hydrochloride is considered a hazardous waste at SERC.

*When finished, always make separate aliquots (20-30uL) of DNA for downstream processing – store these aliquots in the pre-PCR fridge (4ºC; elution buffer dependent) and the stock DNA in the pre-PCR freezer (-20˚C). 

II. DNA QUANTIFICATION

• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILTER (BARIER) PIPETTE TIPS
• This protocol is extremely sensitive and highly prone to cross-contamination. Thus, it is imperative that tubes and bottles not be left open.
• For samples, only open one tube at a time. Do not touch the inside of the tubes with gloves. If you do so, CHANGE GLOVES immediately.
• Additional steps and modifications must be followed during pre-PCR tasks if the
 project targets bacteria, viruses, or human pathogens. Contact Ruth about the training required. 

Follow manufacturer instructions for the NanoDrop2000. Open the software on the desktop. Select Nucleic Acid. Start a new workbook. Name the file “YYYY_MM_DD_NucleicAcid_ProjectName_ExtractionGroupDate.”

a. Example: 2025_11_03_NucleicAcid_SRROyster_ExtractionGroup20251030 

Before starting, mix all samples via vortex, then do a quick spin down on centrifuge to get liquid off the interior lid.

Load 2 µL sample to measure. 

Use the extraction buffer from the kit as the blank. Aliquots are in the drawer next to the NanoDrop. Make fresh as needed. 

Use a Kimwipe to clean the pedestal between samples. 

Record the concentration (ng/uL) and the 260/280 ratio for each sample. Record readings in your lab notebook and/or save results in the CDE Lab folder on the desktop. Create a project folder to save nanodrop results. The results can be copied into an excel spreadsheet and downloaded to a thumb drive to upload to the Team project folder. The computer is not connected to the network. 

III. POLYMERASE CHAIN REACTION (AMPLIFICATION PCR)

*** Repeat 3x each for all samples that are to be included in the metabarcode library ***

• WORK IN PRE-PCR DESIGNATED SPACES ONLY.
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• ALWAYS INCLUDE A POSITIVE AND A NEGATIVE PCR CONTROL
• USE FILTER (BARIER) PIPETTE TIPS
• Mix all samples via vortex, then do quick spin down on centrifuge to get liquid off the interior lid.
• MIX THE FOLLOWING REAGENTS VIA VORTEX: dNTPs, Buffer, MgCl2, primers.
• DO NOT vortex BSA or Taq. Keep Taq cold while in use.
• Only UV-crosslink reagents that DO NOT contain any DNA sequence or enzyme: 10X PCR Buffer, MgCl2, pure H2O
• Additional steps must be followed during pre-PCR tasks if the project targets bacteria, viruses, or human pathogens. Contact Ruth about the training required.

*This protocol is extremely sensitive and highly prone to cross-contamination. Thus, it is imperative that tubes and bottles not be left open.

a. For reagents, remove the lid, pipette out any liquid needed, and place the lid back on top. The lid does not have to be secured in place until it is no longer needed, but it must be covered when not imminently in use.

b. For samples, only open one tube at a time. Do not touch the inside of the tubes with gloves. If you do so, CHANGE GLOVES immediately. 

Use the correct PCR protocol for the primer set for the project. Adjust cocktail mix parameters for the number of samples to be run and volume of template to be used.

Include a negative control (a.k.a., no template control, NTC) and a positive control for each PCR run. The negative control can be crosslinked PCR water (same used to create the cocktail mix) or additional cocktail mix to reach the total reaction volume. Store the positive control tubes in a separate container – DO NOT STORE positive controls in the same containers with sample DNA or PCR reagents.

All users should use aliquoted reagents for dNTPs, BSA, and primers. For stock vials (Gold Buffer, MgCl2, Taq polymerase), note which reagents are currently in use with the date opened (MM/DD/YY) or a distinct mark on the top of the tube. 

Use a primer cocktail for the forward and reverse primers. Primers for metabarcoding are ordered with 0-3 added base pairs. We mix equal volumes of 100µM of each primer iteration into a single primer cocktail for both the forward and reverse primers. For quality control, contact Ruth to create the primer cocktail aliquots. Primer cocktail aliquots for metabarcoding will be labeled with “Ill-All-PrimerName, 100µM” (Ill = Illumina, All = mix of 0-3 added base pairs, primer name for forward or reverse, 100µM concentration). 

Use only UV-crosslinked (2 minutes standard) PCR water (DNAse and RNAse free). The stock bottle must remain in the Coastal Disease Ecology lab, not the Ecological Genomics Core (EGC). Contact Ruth or Calli to create 50mL aliquots as needed for the EGC/CDE to maintain quality control. 

The Taq polymerase must be stored at -20˚C in the pre-PCR freezer when not in use. Use a freezer rack to hold the tube of Taq polymerase during PCR setup or keep it in the freezer until ready to add to the cocktail mix. 

General good practice - record the thermocycler and the start time for the program protocol. Programs are saved in the CDE Lab folder on the thermocyclers. 

After PCR has been performed, reaction products must remain in post-PCR designated spaces. Do not use pre-PCR designated pipettes for post-PCR pipetting. 

Observe proper operation of the pipettes on the PCR bench. If solution is sucked up into the pipette too fast, the pipette itself can become contaminated. It is for this reason that filter tips are incorporated into the workflow. 

IV. GEL ELECTROPHORESIS

*** Repeat 3x each for all samples that are to be included in the metabarcode library ***

• WORK IN POST-PCR DESIGNATED SPACES ONLY
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILTER (BARIER) PIPETTE TIPS
• ALWAYS USE A STANDARD DNA LADDER ON EVERY ROW OF THE GEL
• USE AUTOCLAVE GLOVES TO PROTECT HANDS WHEN HANDLING GLASSWARE THAT HAS BEEN IN THE MICROWAVE
• TO AVOID ELECTROCUTION, TURN OFF THE APPARATUS IF THERE IS ANY SPILLED LIQUID. DO NOT USE DAMAGED RIGS.
• TO AVOID UV DAMAGE TO EYES OR SKIN, DO NOT LOOK DIRECTLY AT UV LIGHT OR DISABLE ANY SAFETY SETTINGS ON THE MULTI-DOC IT 
• Thermometers are available to check the temperature of the melted agarose.
• Mix all samples via vortex, then do quick spin down on centrifuge to get liquid off the interior lid.
• For samples, only open one tube at a time. Do not touch the inside of the tubes with gloves. If you do so, CHANGE GLOVES immediately.

*CONTINUE TO USE FILTER TIPS. At this point, the samples do not have unique bioinformatic barcodes that are used to identify individual samples, so it is imperative that precautions be taken to avoid any cross-contamination.

*Gel electrophoresis is used to confirm that the amplification PCR protocol was successful with the presence of a band on the gel that is the correct size. 

Assemble a gel rig that will accommodate the number of samples that need to be analyzed. One well per row will need to hold DNA ladder. For the X-Large rig (50-well combs), two wells per row should be used (first and last well) as samples on either ends of large gel rigs can move at slightly different rates (a.k.a., smiling). 

If there is no gel in the post-PCR fridge, mix the appropriate amount of agar (tables in box labeled CDE) and clean TBE buffer (in autoclave jars by the gel sink) to generate a 2% w/v agarose gel that will be large enough for all the samples required. Gels can be reused up to three times. 

a. Gel Density calculated as % = weight (g) / volume (mL)

b. We use TopVision Agarose Tablets. Follow the directions on the box for 2% gel.

c. See table below for calculations to mix 2% gel for different sized rigs and how much Gel Red to add. Gel Red is diluted 1:10. Contact Ruth when new aliquots are needed.

Microwave until the agarose has melted. Use an autoclave glove to handle the beaker.

Once the mixture is melted, DO NOT pour into the gel rig until the Erlenmeyer flask is cool enough to hold on the palm of your hand (~52ºC) otherwise you risk permanently warping the gel mold. The XL rig is more sensitive to warping. Cooling the gel to ~48°C works best. Thermometers are available.

Add the appropriate amount of GelRed for the size gel being poured and gently swirl to mix before cooling and pouring. It works best to only add the amount of GelRed you need for a single run. 

Use the bubble level to check that the gel rig setup is level before pouring the mixture into the mold. 

Once poured, allow the gel to cool and become solid (~15 minutes). 

Once cooled, remove the combs and turn the rig so that DNA will move toward the positive electrode. Fill the rig with TBE buffer (from large carboy) so that the agarose gel is completely submerged. Use a disposable dropper pipette to flush any air bubbles out of the wells before loading samples. Dropper can be reused. 

Mix 5uL of PCR product (filter tips only) with 2uL of 6X purple loading dye for each sample on a piece of parafilm. Loading dye is stored in the pre-PCR fridge. Use the loading dye at room temperature (viscous, easier to pipette when at room temperature). Non-filter tips can be used to pipette the loading dye onto parafilm. 

Carefully, load all the samples into the individual wells. You can use non-filter tips for this step. Do not stab through the gel. Do not pipette air bubbles, which will cause the DNA to come out of the well. Make sure there is one well available for the DNA ladder (at beginning or end of row). For the large gel rig (50 samples/comb), load DNA ladder in the first and last wells of each row. For the smaller rigs, only one well for the ladder is required. The 8-multichannel is recommended for loading the XL gel rig 

Briefly vortex mix and spin down the ladder aliquot (NEB Quick-Load Purple DNA Ladder 100bp). Add 10 µL to the appropriate well(s) on each row of samples. 

Put the cover on the rig, plug in the electrodes, and turn on the current. Run at 110 volts for ~35 minutes, depending on the size of the target fragment (running slower at 95 volts for ~45 minutes minimizes “smiling” bands). Check to make sure the loading dye has run far enough down the gel rig before turning off. 

Turn off the electric current, then remove the lid. 

Take a picture of the gel using the Multi-Doc IT. TO AVOID UV DAMAGE TO EYES OR SKIN, DO NOT LOOK DIRECTLY AT UV LIGHT OR DISABLE ANY SAFETY SETTINGS ON THE MULTI-DOC IT. 

Save these pictures in the appropriate project folder (e.g., Teams project, CDE folder on the shared drive). Make sure the image is in focus and not too dark. Save image files in the appropriate Team project folder. 

Name image files with “YYYY-MMDD_ProjectName_PCRType_Maker_PCRTubeIDRange”. 
Example: 2022-05-21_StripedBass_3XPCR_COI_TubesAA12-54. 

Gels can either be broken down, saved and stored in the 4ºC post-PCR fridge (can reuse up to 3X) or thrown away in the general trash flow. When reusing, label the gel flask the date the gel was first made, CDE lab, your name, 2% gel agarose, and “Uses:”. Use tally marks to track the number of times the gel was used. Cover the beaker with parafilm to prevent desiccation. 

V. PCR POOLING

• WORK IN POST-PCR DESIGNATED SPACES ONLY
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILTER (BARIER) PIPETTE TIPS
• Mix all samples via vortex, then do quick spin down on centrifuge to get liquid off the interior lid.
• For samples, only open one tube at a time. Do not touch the inside of the tubes with gloves. If you do so, CHANGE GLOVES immediately. 

Using gel images generated post-electrophoresis, compare the relative brightness of each band for the same replicate across all three replicates. It is highly recommended to use edited gels for this step. 

Based on the comparison across all three replicates, record the required volume to be pooled from each replicate in the project Results Summary excel document.

a. Band brightness generally has three levels: bright, faint, absent.

b. When all bands have the same brightness at a given level (bright, faint, or absent), take 5uL from each replicate:
Ex. R1 = Bright (5uL), R2 = Bright (5uL), R3 = Bright (5uL)
Ex. R1 = absent (5uL), R2 = absent (5uL), R3 = absent (5uL)

c. When all 3 bands are not equal brightness, then take 10uL from the fainter replicates and pool with 5uL from the brighter replicates:
Ex. R1 = faint (5uL), R2 = absent (10uL), R3 = absent (10uL)

d. Note: The proportion pooled is important. Standard is 5uL and 10uL, but other proportions can be used (e.g., 3uL and 6uL) if there is limited product. Further, it is also important that the same proportions assigned to Bright/Faint/Absent are applied to all samples included in the final library. 

Obtain, label, and cross-link (2 minutes) new strip tubes using a letter number combination distinct from the 3XPCR tube labels. 

Using new pipette tips (filter tips only) for each replicate, pipette the triplicate PCR runs for a single sample together so that the appropriate volume from all three PCR runs are combined into one tube. This is now your 3XPCR Pooled product for each of your samples that will be used for indexing. 

VI. INDEXING PCR

• WORK IN POST-PCR DESIGNATED SPACES ONLY
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• DON'T USE TOO MUCH TEMPLATE DNA
• USE FILTER (BARIER) PIPETTE TIPS
• Mix all samples via vortex, then do quick spin down on centrifuge to get liquid off the interior lid. 
*This protocol is extremely sensitive and highly prone to cross-contamination. Thus, it is imperative that tubes and bottles not be left open.

a. For reagents, remove the lid, pipette out any liquid needed, and place the lid back on top. The lid does not have to be secured in place until it is no longer needed, but it must be covered when not imminently in use.

b. For samples, only open one tube at a time. Do not touch the inside of the tubes with gloves. If you do so, CHANGE GLOVES immediately.

Prep work for indexing PCRs must be performed in the post-PCR space even though the bioinformatic barcodes have not been added to distinguish the individual samples. Thus, use caution when mixing. 

Always put on a fresh pair of gloves. 

Make sure each sample is indexed with a unique combination of i5 and i7 primers (dualindexed). There is a template available to help with this step. 

We use KAPA HiFi HotStart reagents for this protocol using the following combination of reagents:

a. Use 1uL of pooled product as template in the indexing PCR. Up to 3uL can be used in cases where project samples have been difficult to amplify. Adjust the cocktail mix calculations accordingly.

b. Indexing PCR done in 25uL total reaction volume using KAPA Ready Mix.

c. 9.5uL water (volume to 25uL), 12.5uL KAPA ReadyMix (1x final concentration), 1uL of 10mM forward and reverse primers (indexing primers – 0.4mM final concentration), 1uL of template (pooled amplicons). 

d. The PCR parameters for indexing depend on how much material you have. Katrina greatly reduced them compared to what individuals do with genomic DNA.

e. PCR water is purchased from Qiagen that is DNAse and RNAse free. To avoid contamination, the stock container must be kept in the Coastal Disease Ecology lab (CDE), not the Ecological Genomics Core (EGC). Aliquots in 50mL tubes are made downstairs in the CDE lab UV clean hood on an as-needed basis to store in the EGC. For quality control, contact Ruth to make new 50mL aliquots from the stock container when needed. Make sure to UV-crosslink (2 minutes) water aliquots before adding additional reagents to the cocktail.

Use strip tubes to create an individual cocktail mix for each i5 and i7 primer. Use the template to calculate reagent volumes for each cocktail mix. 

Use a multi-channel pipette to dispense the cocktail mixes into the appropriate reaction tube. Add 12uL of each i5 and i7 cocktail mix to each reaction tube, following the template to track the combination of barcodes used for each sample (12uL i5 cocktail mix + 12uL i7 cocktail mix + 1uL template = 25uL total reaction). 

Add 1µL DNA template to individual reactions as the last step. 

Vortex mix final strip tubes and spin down briefly to remove liquid from the lids before putting onto the thermocycler. 

Record the thermocycler used and the start time for the program protocol. 

VII. GEL ELECTROPHORESIS

Perform as described above (Part IV) with some changes. 

Spot check that indexing worked by running a gel for a subset of samples, two from each column from the indexing run. Indexing template provides a visual of the concept. 

Do not load control samples. Avoid samples that amplified faintly in the initial triplicate PCRs (if possible). You need to have visible bands on the gel for both pre and post indexed product to determine if indexing was successful. 

Load indexed product and non-indexed product for each sample side-by-side for direct comparison. 

Load 1uL indexed product and 5uL non-indexed product. 

VIII. PURIFICATION WITH KAPA PURE BEADS

• WORK IN POST-PCR DESIGNATED SPACES ONLY
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILER (BARIER) PIPETTE TIPS
• Mix all samples via vortex, then do quick spin down on centrifuge to get liquid off
the interior lid.
• For reagents, remove the lid, pipette out any liquid needed, and place the lid back on top. The lid does not have to be secured in place until it is no longer needed, but it must be covered when not imminently in use.
• DO NOT USE MAGNET IF YOU HAVE A PACE MAKER!!!
• Contact Ruth or Calli for required training for handling hazardous waste. 

Liquid disposed of in this protocol is handled as hazardous waste and must be stored and tracked in an appropriate labeled container in the Satellite Accumulation Area. Contact Ruth and Calli for required training before starting 

Bring both the indexed product and KAPA Pure bead aliquots to room temperature before starting (minimum 30 minutes).

a. Aliquots of KAPA Pure beads are stored in the pre-PCR fridge (2 mL tubes). For quality control, contact Ruth when more aliquots are needed. 

Obtain, label, and cross-link (2 minutes; if not already sterile from packaging) new strip tubes. Close the lids and label tubes using a new letter-number combination that is distinct from the indexing tubes. 

Mix fresh 80% ethanol and cross-link (2 minutes) before using. Use the molecular grade ethanol stored in the EGC flammables cabinet. Use UltraPure water to dilute. For quality control, contact Ruth if more 50mL aliquots of UltraPure water are needed. 

Follow the manufacturer’s protocol for the KAPA Pure beads. Follow for 10uL template input. 

The bead ratio required depends on the target fragment size. Before cleaning, consult with Ruth and Katrina to discuss what ratio will work best for your project. A bead ratio from 0.8X to 1.0X tends to work well for most primer sets regularly used by the CDE lab.
a. Remember that the ratio of beads used depends on the size of the DNA fragment that you are targeting for the project. You can always re-clean your samples to adjust the ratio and remove unwanted small fragments, but you cannot gain back DNA that you lost during cleaning. You may need to clean your samples multiple times. 

Load the DNA template into the strip tubes (one tube per sample; new pipette tip per sample) before adding the KAPA Pure beads. 

Use a clean trough and multi-channel to load the KAPA Pure bead mix to the strip tubes. Use a new pipette tip for each sample. Pipette mix following the manufacturer’s protocol. Only pour out into the trough as much as you need for the reactions you will be cleaning at the time. Put excess beads back into the 2mL aliquot tube using a single channel pipette and new filter tip. 

Magnets are in a labeled drawer in the post-PCR workspace. Two magnet models are available: one to use with strip tubes and one to use with 2 mL centrifuge tubes. DO NOT USE MAGNETS IF YOU HAVE A PACE MAKER!!! 

Do not separate the beads from the final cleaned product. 

IX. QUANTIFICATION WITH QUBIT

• WORK IN POST-PCR DESIGNATED SPACES ONLY
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILTER (BARIER) PIPETTE TIPS
• Mix all samples via vortex, then do quick spin down on centrifuge to get liquid off the interior lid.
• DO NOT USE MAGNET IF YOU HAVE A PACE MAKER!!! 

Bring the samples and Qubit standards to room temperature before starting (30 minutes minimum).

a. Qubit standards are stored at 4°C in the pre-PCR mini fridge.

b. Use the same Lot# standards for the Qubit kit being used. 

Briefly vortex mix and spin down samples. Place onto a magnet until beads have separated and liquid is clear (minimum 2 minutes). DO NOT USE MAGNET IF YOU HAVE A PACE MAKER!!! 

Open only one tube at a time when conducting this protocol to avoid any cross-contamination 

The EGC has two Qubit machines: Qubit and Qubit Flex. Follow the appropriate manufacturer’s protocol for the machine being used.

a. The Qubit measures one sample at a time in microcentrifuge tubes.

b. The Qubit Flex measures 8 samples at a time in strip tubes. This streamlines workflows when processing large numbers of samples (>48). A multichannel can be used to load reagents (from trough) and template (if stored in strip tubes). 

Follow the manufacturer’s instructions for the Qubit High Sensitivity double-stranded DNA kit. 

Use 2 uL DNA template per sample to measure. 

Record sample concentration in ng/uL. 

X. POOLING CALCULATIONS FOR FINAL LIBRARY

Use the equal molar concentration excel template to calculate how much volume of each sample must be added to the final library pool. Libraries (Illumina MiSeq) are limited to 100 – 120 total samples. A separate calculation must be done for each library generated. The template provides additional information about the pooling approach and the lab work.

Have Ruth or Katrina review the final calculation before proceeding with the lab work. Considerations also need to be made if pooling product together from multiple markers with target fragments that differ in size by more than 100bp. Amplicon length variation limits how effectively paired-end sequencing can capture the entire fragment.

Adjust the calculation parameters so that you do not pipette less than 2uL per sample (if possible), as pipettes are less reliable with smaller volumes.

Exclude from the calculation: negative controls and samples whose Qubit concentrations measured “Too Low.”
• Negative Controls: Add 1uL for each sample to the final library pool.
• Too Low: Pool the entire sample volume. For consistency, pool the same volume across these samples. Generally, 30-35uL per sample is available to pool.

Libraries run with a MiSeq v3 600 sequencing kit require >4nM final library concentration. However, higher concentrations are permitted. 

XI. CREATING FINAL LIBRARY POOL

• WORK IN POST-PCR DESIGNATED SPACES ONLY
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILTER (BARIER) PIPETTE TIPS
• Mix all samples via vortex, then do quick spin down on centrifuge to get liquid off the interior lid.
• Open one tube at a time and use a new pipette tip for each sample. 
• DO NOT USE MAGNET IF YOU HAVE A PACE MAKER!!! 

Bring the samples to room temperature before starting (minimum 30 minutes). 

Since KAPA Pure beads are not removed from the product after bead cleaning (Step VIII), use the strip tube magnet to separate beads before pipetting. Briefly vortex mix and spin down the samples. Place onto a magnet to separate the KAPA Pure beads and liquid is clear (minimum 2 minutes).

a. Magnets are in a labeled drawer in the EGC post-PCR workspace. Two magnet models are available: one to use with strip tubes and one to use with 2 mL microcentrifuge tubes.

b. DO NOT USE MAGNET IF YOU HAVE A PACE MAKER!!! 

Label a single microcentrifuge tube with the date (YYYY/MM/DD) name of the library, and “needs final clean”. UV-crosslink this tube (2 minutes). The size tube needed will depend on the estimated final volume in the calculations from Step X. Both 1.5mL and 2.0mL roundbottom tubes are available.

a. NOTE: Each library generated and submitted by the CDE lab (across all projects) is assigned a unique ID prior to submission for sequencing. Contact Ruth to have a unique ID assigned to your final library.

b. Example: L2025E
i. Library = L
ii. Year generated = 2025
iii. 5th library submitted for sequencing in 2025 = E 

Using the equal molar calculations from Step X, pipette the corresponding volume for each sample into the labeled microcentrifuge tube.

a. Print the list of samples and volume to pool from the final library pooling calculation (Step X). Check each sample off the list as it is added to the final pool. 

XII. PURIFICATION OF FINAL LIBRARY WITH KAPA PURE BEADS

• WORK IN POST-PCR DESIGNATED SPACES ONLY
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILTER (BARIER) PIPETTE TIPS 
• Mix all samples via vortex, then do quick spin down on centrifuge to get liquid off the interior lid.
• For reagents, remove the lid, pipette out any liquid needed, and place the lid back on top. The lid does not have to be secured in place until it is no longer needed, but it must be covered when not imminently in use.
• DO NOT USE MAGNET IF YOU HAVE A PACE MAKER!!!
• Liquid disposed of in this protocol is handled as hazardous waste and must be stored and tracked in an appropriate labeled container in the Satellite Accumulation Area. Contact Ruth and Calli for required training before starting. 

Follow the bead cleaning procedure outlined in Step VIII above, with some modifications outlined below. 

Follow the manufacturer’s protocol for the KAPA Pure beads. Use the same bead ratio to clean the final library pool that was used during the initial bead cleaning of individual samples. 

Use 1.5ml or 2.0mL microcentrifuge tubes to perform bead cleaning and to store the final cleaned product to submit for sequencing. UV-crosslink microcentrifuge tubes before using (2 minutes). 

Bring the pooled library that needs cleaning and KAPA Pure bead aliquots to room temperature before starting (minimum 30 minutes). Briefly vortex and spin down both items before using. 

Place the pooled library onto a magnet plate to separate any residual KAPA beads that may have transferred during the final library pooling process in Step XI. Use the magnet plate designed for use with 1.5mL and 2.0mL microcentrifuge tubes. DO NOT USE MAGNET IF YOU HAVE A PACE MAKER!!! 

Bead clean a subset of the pooled library.

a. The subset volume you choose will depend on the initial pooled volume available and estimated molarity from Step X and Step XI. For example, for previous libraries with initial pool volume at 1000uL, a subset of 500uL was cleaned to create the final library to submit for sequencing.

b. If your estimated final molarity is low (< 4nM) from the calculations in step X, bead clean a larger volume and then elute the final library to a smaller volume, thereby concentrating the final library for submission (e.g., bead clean 500uL and elute to 50uL final product).

c. For projects with short target fragments (<300bp) you may need to clean and concentrate the entire pooled library volume available from Step XI to obtain a final molarity >4nM for submission.

Once bead-cleaning is complete, remove the beads and transfer the final cleaned product to a new 1.5 – 2.0 mL tube that is crosslinked and labeled. 

Label the tube with the date “YYYY/MM/DD”, assigned library ID (e.g., L2025E), and “to sequence”. 

XIII. RUNNING TAPESATAON ON FINAL LIBRARY

• WORK IN POST-PCR DESIGNATED SPACES ONLY
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILTER (BARIER) PIPETTE TIPS
• Mix reagents and samples via vortex, and then do quick spin down on centrifuge to get liquid off the interior lid.
• For reagents, remove the lid, pipette out any liquid needed, and place the lid back on top. The lid does not have to be secured in place until it is no longer needed, but it must be covered when not imminently in use. 

Contact Ruth for required training before using the tapestation. 

Follow the manufacturer’s instructions. 

Use the CDE reagents and screentapes that are stored in the pre-PCR mini-fridge (4˚C) 

Bring the reagents, screen tape, and library to room temperature before starting (minimum 30 minutes). 

Tips and tubes can be thrown in the trash when finished. 

Verify the average fragment size for the library is as expected. 

If small fragments are still present (e.g., indexing dimer), bead clean the library again to remove them. Adjusting the bead ratio may be necessary. Re-run tapestation after the second cleaning. 

Once the tapestation results look good, use the equal molar concentration template to calculate the final library molarity (based on the tapestation measurements for fragment size and concentration). 

For LAB library submissions using the MiSeq v3 600 kits, we prepare 50uL of the final, cleaned library at a final concentration of ≥4nM. 

a. You can submit the final library at a higher volume and molarity (it is generally easier for the sequencing techs to dilute a library than to concentrate).

b. NOTE: If you have more than 50uL final product, you can create an aliquot to submit for sequencing instead. Vortex mix well and spin down before transferring the aliquot to a new tube. 

Make sure the sample tube is clearly labeled with the date and assigned Library ID (e.g., L2025E). A printable Word template for cryotags is available on Teams Library Submission channel. Parafilm wrap the final library tube and put into the -20°C post-PCR freezer. Contact Ruth to coordinate sending the library to LAB for sequencing. 

XIV. SUBMITTING THE FINAL LIBRARY FOR SEQUENCING

Parafilm wrap the final library tube and place it into an empty strip tube bag. 

Include a paper label inside the bag. A printable Word template is available on Teams Library Submissions channel. The bag label needs to include:
a. Date (YYYY/MM/DD)
b. Library ID (e.g., L2025E)
c. SERC, CDE Lab
d. Katrina Lohan [email protected]
e. Ruth DiMaria [email protected]
f. Final library pool to sequence 

Store in the post-PCR freezer (-20˚C). 

Contact Ruth and Katrina to provide information needed to submit documentation required for sequencing. A folder with final documentation for the library and submission documentation will be added to the Teams Library Submission channel. 

If shipping libraries, always ship via FedEx overnight, either on dry ice or surrounded by ice packs. Certification is required to ship using dry ice – contact Ruth and Calli. 