Jun 30, 2026
Standard Genetic Methods – Sanger Sequencing Standard Operating Procedures
- Katrina Lohan1,
- Ruth DiMaria1,
- Calli Wise1,
- Emma Palmer1,
- Lael Collins1,
- Tara Sill1
- 1Smithsonian Environmental Research Center, Coastal Disease Ecology Lab
- Coastal Disease Ecology Lab - SERC
Protocol Citation: Katrina Lohan, Ruth DiMaria, Calli Wise, Emma Palmer, Lael Collins, Tara Sill 2026. Standard Genetic Methods – Sanger Sequencing Standard Operating Procedures. protocols.io https://dx.doi.org/10.17504/protocols.io.n2bvj5x2wgk5/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 27, 2026
Last Modified: June 30, 2026
Protocol Integer ID: 319939
Keywords: Sanger Sequencing, sanger sequencing standard operating procedures standard genetic method, sanger sequencing standard operating procedure, standard genetic method, sanger sequencing, 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, Calli Wise, or Katrina Lohan.
I. DNA EXTRACTIONS
• WORK IN PRE-PCR DESIGNATED SPACES ONLY.
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING ASEPTIC TECHNIQUE
• USE FILTER (BARIER) 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.
*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. 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 freezer (-20ºC)
II. DNA QUANTIFICATION
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• USE FILTER (BARIER) PIPETTE TIPS
• 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.”
i. 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)
• 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 mix 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
• For samples and reagents, do not touch the inside of tubes with gloves. If you do so, CHANGE GLOVES immediately.
• Additional steps must be followed during pre-PCR tasks if the project targets bacteria, viruses, or human pathogens. Contact Ruth about the training required.
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 dNTP’s, 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 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 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 pipettors 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
• WORK IN A POST-PCR DESIGNATED SPACE
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• FILTER (BARRIER) PIPETTE TIPS ARE NOT REQUIRED
• 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.
*Unless amplification PCR product is to be sequenced via metabarcoding (if so, reference Metabarcoding SOP), then non-filter pipette tips can be used to transfer DNA into loading dye.
*This step is done to confirm that the amplification 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 PURIFICATION WITH EXO-SAP IT
• WORK IN A POST-PCR DESIGNATED SPACE
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• FILTER (BARRIER) PIPETTE TIPS ARE NOT REQUIRED
• Concentrated EXOSAP-IT is stored in the pre-PCR -20°C freezer.
• Diluted EXOSAP-IT must be stored in the pre-PCR 4°C refrigerator!
• Mix all samples and reagents via vortex, then do quick spin down on centrifuge to get liquid off the interior lid.
Obtain, label, and UV-crosslink (2 min) new strip tubes. One tube per sample. Negative and positive controls are not sequenced.
Stock ExoSAP-IT is stored in the -20°C pre-PCR freezer. Dilutions are stored in the -4°C pre-PCR fridge.
Ensure that you are using diluted (1:10) ExoSAP-IT. To make 1:10 dilution:
a. Label a 1.5mL microcentrifuge tube with “1:10 diluted ExoSAP-IT” and the date it was made (MM/DD/YYYY).
b. UV-crosslink the labeled tube and PCR-grade water for 2 minutes.
c. Briefly vortex the stock ExoSAP-IT tube and spin down.
d. Combine 5uL of stock ExoSAP-IT with 45uL of PCR-grade water.
e. Mix well via vortex, then do a quick spin down on centrifuge to get liquid off the interior lid.
f. Store the dilution aliquot in the pre-PCR 4°C fridge.
For every sample reaction, combine 1uL of diluted ExoSAP-IT with 10uL of PCR product in the newly labeled strip tubes. (You can use less PCR product if for some reason you don’t have that much.)
Run on the thermocycler with the ExoSAP-IT program protocol. Protocols are saved in the CDE Lab folder on the thermocyclers. Record the thermocycler used and the start time for the program protocol.
VI. CYCLE-SEQUENCING PCR
• WORK IN A POST-PCR DESIGNATED SPACE
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• FILTER (BARRIER) PIPETTE TIPS ARE NOT REQUIRED
• Mix all samples and reagents via vortex, then do quick spin down on centrifuge to get liquid off the interior lid.
• Make sure primers are diluted to 10 mM
Use the cycle sequencing template to plan the run before starting.
Obtain, label, and UV-crosslink new strip tubes for 2 minutes.
Create two cocktail mixes – We sequence in both directions, so you need one forward cocktail mix (with only the forward primer) and a reverse cocktail mix (with only the reverse primer).
1X reaction using the v3.1 Big Dye Terminator kit:
a. 6.5 uL of PCR water (UV-crosslinked before adding to cocktail mix)
b. 1.5 uL of 10X sequencing buffer (stored in 4°C pre-PCR fridge)
c. 0.3 uL of 10 mM primer (*one primer per cocktail mix)
d. 0.7 uL of Big Dye (stored in -20°C pre-PCR freezer)***ADD THIS LAST
Leave the BigDye in a cold block or in the freezer while making the cocktail mixes and add it last.
Once set-up is complete, run the SEQ program on the thermocycler. Record the thermocycler used and the start time for the program protocol.
VII. SEQUENCE CLEAN-UP WITH SEPHADEX
• WORK IN A POST-PCR DESIGNATED SPACE
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING AESEPTIC TECHNIQUE
• FILTER (BARRIER) PIPETTE TIPS ARE NOT REQUIRED
• NOTE: Sephadex is a very slippery powder that is also very expensive, so we keep it in a plastic container to avoid spillage. Change gloves once finished handling the Sephadex as your gloves will be slippery.
This step removes all remaining primers and dideoxy nucleotides from the cycle sequencing products, which can cause problems during sequencing and generate messy sequence data.
Follow instructions provided in Sephadex Protocol 2026.
Note: LAB requires semi-skirt plates to submit samples for Sanger sequencing. Check sequencing lab requirements if other companies are to be used for your project.
Millipore plates are reused. Run DI water through the plates via centrifugation prior to adding and hydrating Sephadex. This ensures that the wells are clean and minimizes failure of samples to spin through the hydrated Sephadex.
If you hydrate Sephadex overnight, place the plate with a paper towel moistened with DI water into a plastic bag and put it into the -4°C post-PCR fridge to prevent desiccation. Given the sensitivity of spinning samples through hydrated Sephadex, CDE generally hydrates and spins samples through Sephadex on the same day.
Once finished, seal the labeled submission plate with foil and place it in the -20°C postPCR freezer.
VIII. STANDARD RECIPIES AND STOCKS
*** All stocks must be made with fresh aliquots of PCR-grade pure water ***
Primer Hydration:
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING ASEPTIC TECHNIQUE
• ONLY USE FILTER (BARIER) PIPETTE TIPS
• ONLY WORK IN PRE-PCR CLEAN BENCHES
• USE FRESH ALIQUOTS OF PCR-GRADE PURE WATER (DNAse/RNAse FREE) FROM THE STOCK BOTTLE STORED IN THE CDE LAB
• 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.
• Contact Ruth or Calli to assist with making aliquots to maintain quality control.
• Additional steps must be followed during pre-PCR tasks if the project targets bacteria, viruses, or human pathogens. Contact Ruth about the training required.
When primers are ordered from a company, they arrive in a dry state (referred to as “dried down”) and must be reconstituted before use to 100mM.
To create 100mM primer stocks, you must add the appropriate amount of PCR-grade pure water (RNAse and DNAse free) to each tube. The volume of water will differ for each primer.
Remove 50-100L aliquots for individual use. Do not freeze-thaw the stock each time as degradation will eventually occur. A conventional 25L PCR generally uses 0.5L of each primer.
Before beginning, label the top of each stock tube with the primer name and “100mM”
Using a personal centrifuge, centrifuge all the tubes for ~5 seconds. This will ensure that all the dried-down primers are located at the bottom of the tubes.
Water used for hydration MUST be clean!!! UV-crosslink the PCR-grade water specifically set aside for primer hydration. You MUST use filter tips and a new tip for each tube containing primer.
a. NOTE: For the UV crosslinker to be fully effective, plastics and reagents should be opened before being placed into the machine. Screw caps should be placed with the inside of the cap facing up. Close all tubes and reagents immediately upon removal from the machine.
b. DO NOT put the primers in the crosslinker – they will be destroyed!
On the individual primer stock tubes, the nmol amount is listed. You must add 10X the amount of water as the nmol listed to create a 100mM stock.
a. Example: if a primer is listed as being 18.8 nmol, then you add 188 µL of water to create 100 mM stock
Once the water is added, vortex at maximum speed for ~5 seconds to homogenize the solution
Aliquot 20-30 µL into a 0.5 mL tube, which should be UV crosslinked before filling and labeled with the primer name and “100mM” solution. Dilutions of primers should be made from the working aliquots, not the freezer stock tube.
Store working primer aliquots at 4ºC (project specific) and primer stocks at -20ºC.
Primer Aliquoting:
*** For amplification PCR we use 100 mM primers ***
*** For cycle sequencing PCR we use 10 mM primers ***
• WEAR CLEAN GLOVES
• WORK CAREFULLY USING ASEPTIC TECHNIQUE
• ONLY USE FILTER (BARIER) PIPETTE TIPS
• WORK UNDER PRE-PCR CLEAN BENCHES
• 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.
• Contact Ruth or Calli to assist with making aliquots to maintain quality control.
• Additional steps must be followed during pre-PCR tasks if the project targets bacteria, viruses, or human pathogens. Contact Ruth about the training required.
To aliquot primers, get a new 0.5 mL tube. Label the top of the tube with the name of the primer and “100mM.” Put it in the UV-crosslinker for 2 minutes.
a. NOTE: For the UV crosslinker to be fully effective, plastics and reagents should be opened before being placed into the machine. Screw caps should be placed with the inside of the cap facing up. Close all tubes and reagents immediately upon removal from the machine.
b. DO NOT put the primers in the crosslinker – they will be destroyed!
In the clean hood, thaw the primer stock solution. The tube must be completely thawed before an aliquot is taken.
Vortex the thawed solution and then quickly spin down the in the personal centrifuge to remove liquid from the sides and inside the cap.
Using filter tips, pipette 20-30µL of primer solution into the clean 0.5mL tube.
Return the stock to the -20ºC pre-PCR freezer. Store the primer aliquot at 4ºC (project dependent).
Create dilutions from the 100 mM primer working aliquots as needed for projects.