Kartzinel:Laboratory Protocols

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All lab users must complete training requirements and consult with PI Tyler Kartzinel before beginning research in the lab. This wiki is also required reading. Please refer back to it regularly because policies and protocols are subject to change as we improve our work. If details are missing, do not assume they are unwanted; consult with Tyler to identify appropriate protocols and remember to report successes for inclusion on the wiki.

Pre- and Post-PCR

The lab will adhere to strict pre- and post-PCR policies. The general flow of materials through the DNA lab will be sample organization and extraction (pre-PCR freezer room and Biosafety cabinet), PCR setup (PCR hood), and post-PCR processes (thermocycler, gel rig, post-PCR freezer).

  • Samples, extract, and pre-PCR reagents must be kept in the pre-PCR room.
  • PCR reactions will be set up in the pre-PCR room, then moved to the thermocycler in the post-PCR room. All post-PCR applications must happen in the post-PCR room.
  • There is a designated set of sample racks for sample organization and extraction (green or blue), PCR setup (pink or purple), transfer (orange), and post-PCR applications (yellow).
  • Samples, reagents, and materials that need to move from the pre-PCR room to the post-PCR room have to happen in transfer containers. These are racks, coolers, and tubs that will NEVER BE USED WITH OPEN SAMPLES in either room. The sole purpose of the transfer vehicles is to physically move your stuff from one room to the other. They should never be used around pre-PCR samples or reagents (to avoid accidentally introducing post-PCR products) and they should never be used around post-PCR samples or reagents (to avoid accidentally picking up post-PCR products). Transfer containers are available for PCR strip tubes/plates, Qubit assay tubes, milliQ water, etc.
  • The transfer station in the pre-PCR room is designated next to the emergency eyewash station, to avoid accumulating post-PCR aerosols. Ensure the transfer racks are stored in their container. Keep container closed. Surface-sterilize transfer racks with bleach immediately upon returning them to the pre-PCR room.
  • The transfer station in the post-PCR room is designated on the metal rack near the door.

General Guidelines

Cleaning lab resources

  • All lab users are responsible for the cleaning and maintenance of each piece of lab equipment each time they use it (see equipment-specific details below).
  • All lab users are responsible for cleaning and storing the shared tube racks and trays that they used. If cleaning requires soaking something, please do not forget to return and finish the clean up as soon as appropriate.

Cleaning solutions

Note that bleach solution destroys both living cells and free nucleotides (contaminant DNA). Denatured ethanol will also destroy living cells, but is less effective at removing contaminant DNA. Therefore, we use bleach as a primary means of cleaning and decontaminating. The ethanol solution is useful for rinsing bleach from surfaces and also evaporates quickly, which is important because we don't want bleach carrying over onto your samples or clothes. Ethanol solution is also useful for decontaminating goggles.
Making solutions

  • 10% Bleach: Make a fresh solution daily in the bleach-specific squirt bottles. Pour about 5 mL bleach and fill to 50 mL (i.e., adding 45 mL) with tap water (measure with beakers/graduated cylinders). Avoid waste: if you typically make more than you use, scale back these volumes!
  • 20% Bleach: Make a fresh solution daily in the bleach-specific squirt bottles. Pour about 10 mL bleach and fill to 50 mL (i.e., adding 40 mL) with tap water (measure with beakers/graduated cylinders). Avoid waste: if you typically make more than you use, scale back these volumes!
  • 70% Ethanol (denatured): Use only denatured (cheap) ethanol for cleaning. Pour about 30 mL denatured ethanol into the ethanol-specific squirt bottle and fill to 100 mL. This bottle should be dated using lab tape and alcohol-resistant sharpie. This solution does not need to be made fresh daily, but alcohol evaporates and should be replenished every couple weeks.

How to clean bench and hood surfaces

  • Bleach requires 15 min contact time to work. Squirt bleach on the bench, hood, or compatible equipment surface; wipe it around using paper towel or kimwipe and set a 15-min timer (use paper towel for most bench surfaces; kimwipe for more sensitive surfaces where noted below). Then rinse with ethanol solution and paper towel or kimwipe.

How to decontaminate biohazardous liquid waste

  • As liquid waste accumulates at the bench, add approximately equal volumes of your 20% bleach solution -- or approximately 10% volume of fresh bleach -- to achieve a final concentration of 10% bleach. The decontaminated solution should then be moved to the satellite liquid waste accumulation containers.


  • To wash pre-PCR tube racks and trays, use 10% bleach. Use squirt bottle and paper towel to make sure surfaces have contact. Allow 15 min of contact time. Rinse.
  • To wash large numbers of racks and trays, fill a wash basin with 10% bleach. Make sure items are fully submerged (no air bubbles) and that anything sticky is scrubbed off. After at least 15 min contact time rinse with tap water, shake out, and allow to dry on the racks. When fully dry, make sure these are put away in their appropriate (color-coded) cabinets. If you leave them out, they will become dirty through disuse, and your colleagues (or you!) will have to wash them a second time.


  • To wash post-PCR tube racks, follow the same protocol as pre-PCR tube racks, but keep everything in the post-PCR room.
  • To wash post-PCR flasks used to pour gels, scrub them out with warm water while still warm (before agarose cools and solidifies). Check interior for residual agarose.

Stock Materials

  • We maintain labelled "inventory containers" under the bench across the from the biosafety cabinet in the main lab and added to the bottom wire shelf in the post-PCR lab. This allows us to resupply shared resources before they are depleted.
  • When stock reagent bottles or supplies boxes are empty, please place them in their respective "inventory containers" in the lab.
  • For supplies that come in boxes like gloves, tips, and tubes, please remove the side of the box with the product information (tip volume, glove size etc.) to save space.
  • The Molecular Lab Leader checks inventory containers approximately weekly to initiate resupplies. If you plan to use an unusually large quantity of supplies in a short period of time, please coordinate with the PI to ensure our resupply strategy can keep up.

Equipment Use & Maintenance

It is each lab member's responsibility to read equipment manuals. Check with the PI for input.

Rees Freezer Alert Systems

  • The -80 and -20 freezers in the main lab, which are used for sample and reagent storage, are programed to a temperature alert system.
  • Lab members will receive alerts by phone and email if the temperatures rise above (or fall below) a set point.
  • If you accidentally keep the door open long enough to trigger this alert, please close the door and wait for the unit to get back to temperature; send a reassuring email to the lab group.
  • Otherwise, it is critical that all lab members who receive the alert coordinate a prompt response (someone will have to check the unit and respond appropriately).
  • We have access to emergency backup freezers if necessary; the PI's emergency number is posted on the lab door (call/text day or night if an urgent response is needed).
  • In case of malfunction or maintenance needs, more information on the Rees system can be found here: https://www.brown.edu/about/administration/biomed/facilities-planning-operations/research-operations/rees-monitoring-system.

-80 Freezer

  • Keep the freezer locked. Do not leave the key in the door when you exit the freezer room.
  • Close the door, latch the handle, and lock the door in three separate motions to ensure a good seal. You may need to press gently on the front of the external door door and ensure internal doors are closed to ensure it latches properly.
  • Press the pressure release button (bottom left side near the door) occasionally to avoid build up of frost.
  • If the door is sealed due to pressure build up, press the pressure release button.

Manual pipettes

Pipettes are vital to the quality of your work, and they are expensive. All users should adopt our general guidelines.

  • Use only filter tips for pipettes (only filter tips should be ordered for the lab).
  • Place them gently in their holders when not in use; return them to exactly their maximum setting when not in use (e.g., the P-200 should be set to 200 μL).
  • Place them in their holders rather than on the bench while working in order to keep them clean and out of danger of spills/contamination.
  • Use pipettes for volumes near the center of their limit (e.g., do not use a 20-200 μL pipette to pipette 200 μL; use a 100-1000 μL pipette instead).
  • If you notice a pipette is not working smoothly, pause your work and report it or ask for help cleaning it.
  • Remember to close the lid to the pipette tip boxes to keep them clean and secure!

Repeating Pipette

  • Use requires training from a member of the lab with prior experience.
  • Never force tips in or out of the unit. Do not twist tips when inserting or removing them from the unit.
  • Most of the time, we will use the "dis" setting, which withdraws liquid and then aliquots a specified amount repeatedly.
  • Use only for making aliquots (e.g., preparing lysis tubes for the field; dispensing molecular grade water into 1.7 mL tubes).
  • Plug in after use to ensure the unit is charged for the next user.

Biosafety Cabinet

  • This cabinet should only be used for sample manipulation and DNA extraction. No PCR reagents or PCR products should be put into this cabinet.
  • Never assume the person before you did a good job cleaning and stowing the cabinet: use freshly prepared 10% bleach (<24 hours old) to clean surfaces and equipment (and rinse with cleaning ethanol). Run the hood for 15-30 minutes with the UV light.
  • Always clean and stow the cabinet after use: properly secure solid/liquid waste, hang up pipettes, turn off equipment, use 10% bleach to clean surfaces and equipment (and rinse with cleaning ethanol), run the hood for 15-30 minutes with the UV light, and then finally turn off the hood.
  • Remove user- and project-specific items from the cabinet (e.g., bags of tubes).

PCR hood

  • The only things that should happen inside this hood are related to PCR set up (including aliquoting and diluting of PCR stock reagents).
  • Stock reagents kept at room temperature (molecular water, DMSO) are kept in a cabinet over the main bench and behind the hood. Consult with the PI about project-specific aliquots for your work.
  • Racks, trays, and tubes for PCR setup are kept in a glass cabinet to the right of the PCR hood. These must be bleached and rinsed between uses, and should be returned to the cabinet in this condition.
  • Daily stocks of tubes, strips, and plates can be found in buckets and bags at the hood (shelves above and drawers beside).
  • Long-term stocks of tubes, strips, and plates can be found in labeled shelves and cabinets, mainly along the office wall. These can re-stock the 'daily-use' supplies at the hood.
  • 15 min prior to use, use 10% bleach to wipe down the interior of the cabinet and wipe down with paper towels. Rinse with 70% denatured ethanol. Run the UV. Tube racks and bags intended for use can be added at this time to be bathed in UV.
  • Make use of the refrigerator and freezer at the bench. These are for short-term holding of reagents and samples. You can bring reagents and samples out of long-term (alarm-monitored) freezers for PCR setup(s).
  • When setting up reactions or aliquots: run the UV recirculator fan constantly, and close the sash whenever possible if walking away or pausing in the setup. When closing the sash, you should run the UV whenever possible, but NEVER close the sash and allow the UV to turn on when samples/reagents are inside; practice first to ensure you have the correct settings.
  • Raw samples and PCR products should never be brought into this hood.
  • All stock and working stock reagents should be put away before introducing DNA samples to the hood. Only after reagents are put away should DNA be aliquoted to reaction tubes. Make good use of the bench-side freezer and refrigerator for efficiency.
  • When finished, pipettes, equipment, and open tip-boxes can remain inside, but do not store bags of tubes, strips, etc. inside.
  • To clean after use, use 10% bleach to wipe down the interior of the cabinet and wipe down with paper towels. Rinse with 70% denatured ethanol. Run the UV floodlights on the 15-min timer.
  • Remove user- and project-specific items from the cabinet (e.g., bags of tubes).
  • Samples and reagents can remain in bench-side refrigerator and freezer for short periods of time (days-weeks) while projects are ongoing, but should be moved to long-term storage otherwise.

Bench side refrigerator and freezer

  • There is a -20°C "mini" freezer and 4°C refrigerator at the PCR station to facilitate PCR setup.
  • This freezer is not on a backup outlet, and it is not connected to a temperature alert system. Therefore, it is not safe for long-term storage of samples or reagents.
  • User stocks of reagents, including primers and taq, can be placed in this freezer before and after PCR setup to minimize back-and-forth trips to the freezer room as well as the time tubes are kept at room temperature.
  • Lab stocks should not be kept in this freezer. Long-term sample storage should not rely on this freezer.
  • Please check the mercury-free freezer thermometer when adding/removing tubes from this freezer, in case adjustments are required.


  • Store reagent and buffer at room temperature on the shelf, in the original box so that the reagent is not exposed to light. Store standards in the refrigerator at 4°C.
  • We use the high-sensitivity assay for most applications in the lab; a broad-range assay can be used when appropriate. Refer to manuals and consult the PI.
  • Prepare master mix to include 2 μL of sample or PCR product; for most of our work, 1 μL could yield less precision and >2 μL could be wasteful.
  • Prepare master mixes of 1000 μL for batches comprising a total of 10+ samples and standards. Less is fine, if <10 samples and standards will be used. Do not use very large tubes to prepare mixes for 10+ samples and standards (it will be awkward with our pipette sizes, and precision will suffer).
  • Use a vortex and minicentrifuge to effectively mix your master mix prior to aliquoting to tubes, as well as your assay tubes after adding the sample/standard.

Gel Rig

The gel rig must be cleaned after each use. The TBE buffer can be reused a limited number of times, but must be stored separately in an air-tight container. Buffer cannot be stored in the rig itself because this will lead to deterioration and it will alter buffering capacity through evaporation. The rig and lid can be rinsed with warm water, checked for scraps of agarose that need to be removed, and be inverted to dry. Do not wash the rig with bleach.

Gel Imager

Our gel protocols are available below. The imager is compatible with a USB memory drive. Everyone should save copies of their images using the USB, and upload them to LabArchives. Gels must be removed and the transluminator surface must be cleaned with Kimwipes (NOT paper towels) after each use. To remove sticky agarose, try dampening a kimwipe with a bit of warm tap water. Use only clean gloves on the exterior and clean any inadvertent buffer/gel from the exterior with a bit of warm water and a clean kimwipe (you and our colleagues will be plugging USB sticks into this devise, and wouldn't touch those with dirty gloves).

Reagent storage and handling

Room temperature stocks

  • There are three places to look for shared stock items: (a) the flammables cabinet for reagents such as EtOH and (b) a cabinet above the main bench for reagents such as molecular grade water, DMSO, trehalose, etc.
  • Individuals should take aliquots for specific projects, but must adhere to key policies: (a) ensure a re-order is approved when stocks run low and (b) calculate the volume needed in advance to minimize wasted resources.

Freezer stocks

  • Original primer stocks (100 μM) are kept in a box in the -80°C for long-term storage.
  • Working stock primers (10 μM) are kept in the -20°C.
  • Polymerases, BSA, and dNTPs are kept in cold blocks in the -20°C.
  • Post-PCR sequencing primers (1 μM) are kept in the post-PCR -20°C.


  • Common enzymes include all polymerases (taqs), BSA...
  • Store enzymes at appropriate temperature (based on product inserts). Taqs and BSA should be kept in the -20 inside the freezer block dedicated exclusively to reagents (no samples!), to minimize temperature fluctuations.
  • NEVER vortex or centrifuge enzyme reagent vials; master mixes containing these enzymes, such as PCR recipes, should be mixed and spun prior to starting the reaction.
  • Temperature-sensitive enzymes should be left frozen until the last minute, and quickly returned to storage. When setting up a PCR reaction, prepare the entire master mix before taking taq/BSA out of the freezer, and quickly return it. To facilitate, we have a bench-side -20°C freezer and a portable cold block to store and help transfer reagents between short- and long-term freezers. In other words, we have two portable bench top coolers are available, one for transporting your reagents from the freezer to the PCR hood and another to hold your master mix while preparing reactions. The cooler holding your master mix should be cleaned after each use.

Temperature sensitives

  • Temperature sensitive reagents that are not enzymes include Qubit standards, Qubit reagents, dNTPs, primers, beta-mercaptoethanol, gel ladders ...

Light sensitives

  • Common light-sensitive reagents include Qubit reagents and gel dyes.
  • Light-sensitive reagents should be left in their storage box until use, and returned promptly.

Resuspending primers

  • Before resuspending primers, clean the PCR cabinet and obtain a fresh aliquot of molecular grade water.
  • Record your initials and the date on the tubes.
  • Spin primer stocks in the mini centrifuge for about 10 s before opening (dry primers are a powdery residue that may get stuck around cap during shipping).
  • Prepare 100 μM stocks by adding X μL molecular grade water, where X = 10 * the nmol of primer listed on the side of the tube.
  • Vortex the primer stocks and give them time to resuspend before use in PCR. This could be an hour at room temperature, or overnight in the refrigerator at 4°C, in order to ensure proper molarity. (Plan ahead.)
  • Make ~5 aliquots of 10 μM working stocks at 100 μL. Store the working stock aliquots in the -20°C and the original stocks in the -80°C.
  • Note that "Lab Ready" primers do not require this preparation before use, as we receive them at 100 μM concentration.

Waste Disposal

  • All chemicals (including boxes of zymo reagents) have a Brown University chemical inventory barcode on them. When empty, please remove the barcode and stick it to collection paper (taped to the side of the fume hood above the red liquid-waste collection bin).
  • Work with certain samples--especially those of international origin--require additional waste disposal considerations. See our lab's Biosafety Authorization or talk to the PI for more details.

Liquid waste

Liquid wastes should be placed in the container labeled with an orange hazardous waste sticker, and put in secondary containment (red plastic basins). Both pre- and post- PCR we have designated sites for liquid waste.

Solid waste

Solid waste that is in contact with samples should be placed in a sealed plastic bag and placed in a red bag lined box. These boxes are available in the pre- and post- PCR rooms. Do not place liquids in these boxes.

Freezer Stocks


Most primers in the lab fall into one of two categories: (i) plain old locus-specific primers and (ii) tailed primers engineered for preparing Illumina libraries using Nextera/Nextera-like protocols.

Current stocks of locus-specific primers can be found HERE.

  • The google doc is viewable by anyone with a Brown University email address; contact TRK for access/edit permissions.
  • Primers are listed and stored in the freezer together in pairs; sometimes 'pairs' of primers can be used interchangeably, creating options for tailoring based on taxon or target sequence-length.
  • Protocols for typical PCRs can be found listed in the section on PCR, below. (Add link).
  • Please keep track of when initial stocks were ordered ('lab ready' or rehydrated), initial stock volume, and the working-stock aliquots remaining.

Current stocks of tailed primers for Illumina sequencing can be found HERE.

  • The google doc is viewable by anyone with a Brown University email address; contact TRK for access/edit permissions.
  • Primers are listed and stored in the freezer together in pairs; sometimes 'pairs' of primers can be used interchangeably, creating options for tailoring based on taxon or target sequence-length.
  • Protocols for typical PCRs can be found listed in the section on PCR, below. (Add link).
  • Please keep track of when initial stocks were ordered ('lab ready' or rehydrated), initial stock volume, and the working-stock aliquots remaining.


Current reagent stocks and uses can be found HERE.

  • The google doc is viewable by anyone with a Brown University email address; contact TRK for access/edit permissions.
  • As much as possible, we aim to economize and simplify lab work by developing shared protocols. This way we can order in bulk, learn from each other, and keep organized. This does not mean we are inflexible in the lab, as individual projects may call for more specialized resources. So, we list reagents for 'standard' and 'misc' protocols on separate tabs.
  • The set of 'Misc' reagents include (i) those for specialized (or expensive) applications that we aim to minimize use of and (ii) those leftover from projects using 'old' protocols that we don't use much anymore.
  • Some 'standard' reagents are being 'phased out' of the lab: we continue to use them for ongoing projects to maintain consistency (e.g., some long-term metabarcoding projects need to use the same polymerase we began using years ago), but should not be used for new projects.

DNA sampling & extraction protocols

Plant DNA Barcode Reference Library

We are often asked for guidance and assistance building plant DNA reference libraries. The strategies and logistics involved will inevitably differ depending on the goals of a research project, conditions in the field, and access to resources such as existing herbarium collections and experts on the local flora. Here we provide some general suggestions based on experience. Our general Protocol for Collecting Plant Vouchers and DNA Samples is available HERE.

  • The google doc is viewable by anyone with a Brown University email address; contact TRK for access/edit permissions.
  • The doc refers to two metadata sheets. One for is collecting herbarium voucher specimens that can be submitted for mounting, archiving, and digitization at Brown University; this can be requested from the Director of the Brown Herbarium or by emailing TRK. The second is for collecting plant DNA barcode samples that is compatible with the barcode of life data systems, which can be downloaded from boldsystems.org.

Zymo Soil & Fecal Mini Kits

This is the kit we use for most of our fecal DNA research. Please note that the company can change (and has historically refined) the protocol. For this reason, the wiki does not recapitulate information contained within the protocol. You must refer to the protocol from the box you are using to ensure you have the correct version. We simply provide helpful tips for organizing and maintaining the highest standards for your workflow.

Preservation and Lysis Buffers

  • Standard Buffer is fine to use for samples that are obtained frozen, dried, or on EtOH. It is cheap and suitable.
  • Xpedition Buffer (antiquated) and DNA/RNA Shield Buffer (more recent) are designed to lyse and preserve samples in the field. They are more expensive because they need to be ordered separately. See notes on preparation below.

Preparing Lysis Tubes Prior to Sampling

  • We inventory and prepare sample lysis tubes prior to beginning fieldwork whenever possible.
  • Prepare tubes in the biosafety cabinet after proper preparation, which involves wiping all surfaces and equipment inside the cabinet with 10% bleach and running the UV light for 30 minutes (a kitchen timer is provided).
  • Use the repeating pipette to aliquot the required amount of buffer into your tubes.
  • A piece of aluminum foil (internal side up) can be placed on the workbench inside the cabinet to provide a clean surface where you can place your tube caps efficiently (internal side down to prevent aerosols and dust from settling inside).
  • When re-capping, really crank the caps back on because the O-rings have allowed leaks in the past. Be aware and discard tubes that fail to seal properly. Report this to Tyler.
  • Apply pre-labeled stickers with appropriately procured Quartzy inventory numbers. Stickers are available in the lab or from Fisher Scientific (Cat. No. 15930C). They must be laser printed.
  • Randomly select at least one tube per box to serve as the "blank" for the batch. Use sharpie to write the word "BLANK" in addition to the inventory number to avoid inadvertently filling it with a sample in the field. The blank field should also be included in the inventory number on Quartzy (‘TK####### (Extraction Blank)’).
  • Prefilled and labeled tubes can go back in the baggie for use in the field. Write your initials, the range of inventory numbers included in the bag, date filled, and batch number on the exterior of the bag. Make sure the same is written on the box containing the rest of the reagents and columns. This box can stay in the lab (for use when completing extractions).

Field Sampling Considerations

  • For many lab members, it will be possible to develop a research plan based on existing protocols. Specific protocols and standard operating procedures that have been used and approved in the lab can be found in our Lab Archives account and/or by consulting with Tyler. These protocols include details pertaining to sample handling, preservation, transport, permitting, etc.
  • Considerations for effective lysis in the field, when this is the appropriate preservation method: (a) this is perhaps the most critical step in ensuring high-quality DNA is recovered, (b) use the most powerful vortex available, ideally horizontal on a wide top or in clamps, (c) run the vortex for at least 30 s, but approximately 2 min can lead to better quality results (pause every ~30 s to ensure the vortex motor does not overheat if you are not using a unit with this builtin safety feature), (d) check visually for complete homogenization of the sample (if you see chunks, continue vortexing for up to ~5 min; longer may result in unwanted shearing of the DNA and is not generally a good solution compared to making adjustments in starting material composition -- use less material and smash it up prior to loading the tube -- remember that more lysis buffer can be added in the lab if needed).

Please Note: Mini vs. Micro kits There are zymo "micro" kits in the lab. Be aware of the differences between the mini and micro kit. We find the mini kit works very nicely for most of our fecal DNA analyses, but the micro kit is problematic. The micro kit columns are especially prone to clogging, and therefore should only be used in projects involving very small quantities of input material. If clogging occurs using while using the Zymo-Spin II-F Filter (red top filter) in the micro protocol, use the filtrate and move on to the next step. DO NOT CENTRIFUGE MORE THAN ONCE. If clogging occurs using filters at any other stage centrifuge until ALL liquid passes through the filter. THIS MAY REQUIRE MULTIPLE ROUNDS OF CENTRIFUGING.

Preparing for Zymo extractions

  • Please use beta mercaptoethanol in buffers. Because this is not a long-term shelf stable mixture, please add just before using the kit for the first time and try to use up the entire kit within a week or so.
  • We store the stock beta mercaptoethanol @ 4 degrees.

Plant Extraction Kits

Qiagen DNAEasy Plant Mini The kit that RYK uses

Blood and Tissue

Qiagen DNAEasy Mini

  • FTE cards and biopsy punches...
  • Cleaning biopsy punches...

PCR protocols


PCR introduction

A PCR contains the following necessary reagents:

  • PCR-buffer. Salt and pH-stabiliser. User stock of 10x can be kept in your box.
  • MgCl2. Salt which is required for the polymerase to work. Standard reaction concentration is 1.5 mM (ranges from 1-4 and can be optimized). Higher concentrations make polymerase less specific and favor amplifications of short fragments. Too much MgCl2 often results in multiple non-specific bands. An aliquot of the stock can be kept in your personal PCR box.
  • dNTPs. Free nucleotides (Gs, As, Ts and Cs) that are strung together to make DNA copies. An aliquot of the user stock (10 mM of dNTP Mix, which has 2.5 mM of each dNTP) can be kept in your personal PCR box.
  • Primers. Single stranded DNA (oligonucleotides), usually 18-30 bp. Stock solutions are kept at 100 μM, and user stocks are diluted to 10µM for use and storage in your box.
  • Polymerase. The enzyme that strings nucleotides together. It starts at the 3' end of the primer and uses the complementary DNA strand as a template. User stock of X units/μl is kept in the freezer block to ensure constant cold storage.
  • Template DNA. The source of DNA for the PCR amplification (i.e., your sample). We assume a standard concentration of 25 ng/μl, but depending on the organism, sample quality, and extraction protocol you could have a big range of concentrations (not uncommon for us to have (1-100 ng/μl). If your project has a large range of concentrations, or variable amplification success, template concentration can be quantified (by Qubit) and quality assessed (with a gel run) in the lab to help improve consistency.

In addition, some reagents can improve PCR success if needed:

  • BSA (bovine serum albumin). An enzyme that prevents binding of DNA to the reaction tube. It is recommended for use at 10-100 μg/ml.
  • DMSO (dimethylsulfide). A chemical that reduces formation of secondary DNA structures (i.e., it linearizes DNA). It is recommended for use at 1-10% DMSO.
  • Master mix. PCR recipes are reported on a per-reaction (i.e., 1X) basis, and we calculate a "master mix" based on this recipe. The master mix is simply the 1x recipe multiplied by the number of reactions that will be set up, including all positive and negative controls (see below). The number of reactions used to calculate the master mix recipe should be the number of reactions + 10%. The master mix contains all components of your PCR recipe except the template. Components should be added in order of decreasing volume and expense (so water should be added first, and polymerase last). User stock reagents should be vortexed and spun down to ensure homogeneous mixtures and minimize risk of contamination. However, enzymes (BSA, polymerase) should NOT be vortexed or spun, and this is one reason why they must be stored upright in the cold block. Once the mix is complete (including polymerase), it should be vortexed and spun before aliquoting to reaction tubes (where the template will be added).
  • Fancy pants. Some protocols use pre-mixed PCR reagents. Just add primers and template. We use these sometimes. They can be wildly expensive and we should not default to them when other protocols may be suitable.

PCR controls

  • Negative control. Also known as "no template control" or "water blank". Every single PCR setup needs to include at least one negative control. The negative control is simply a reaction that includes molecular grade water (the same user stock of water used to make your master mix!), added with the same volume and in place of the template DNA.
  • Postitive control. Positive controls should be run whenever possible. Some new protocols may need to be piloted without an obvious positive control available for use, but once a protocol is known to be working it is helpful to include a standard sample as a positive control to help detect inconsistencies. This can be a "mock community" (common in microbiome studies) or an unimportant sample of similar genetic composition to your target and that is known to work with your protocol (e.g., tree DNA extracted from a plant on campus, as a high-quality DNA source for comparison to plant DNA contained within herbivore dung).

PCR setup

  • All PCR reactions should be set up in the PCR hood (see comments above for rules about using and maintaining the hood).
  • Please note that we use "cold blocks" rather than wet ice to keep reagents, samples, and mixes cold during setup. We do not use wet ice because it increases risk of contamination (condensation, dripping ice, unstable tubes). If ever necessary due to a shortage of cold blocks, wet ice can be used, but tubes should NEVER be placed directly in the ice. Instead, place a tube rack into the ice bucket or use a cold block designed to be filled by wet ice to ensure stability.
  • Put a ziplock/empty bag in the tip bucket. This is useful for absorbing aerosols when ejecting tips, keeping the workspace clean, and disposing of tips/tubes in the red box. Do not allow this bag to become top heavy or more than 2/3 full, to ensure waste is kept away from reagents and samples.
  • Organize the hood ergonomically, aiming to minimize long-distance motions and carry-overs (i.e., prevent hands, dirty pipette tips, sample tubes, etc. from being carried or opened directly above other samples, reagents, or reaction tubes. If right-handed, keep the pipette carousel on the right and within easy reach so you can grab and hang up each pipette easily; keep tip boxes just in front of the carousel for easy loading; keep the trash bucket in easy reach (to avoid "shooting" tips towards it, and behind the tip boxes to keep it away from samples/reagents; keep the vortex and mini-centrifuge to back left by the outlets; keep reagents/samples in front of that; keep a clean and open space for aliquoting reagents and samples in the center.
  • Once the bench is cleaned and ready to go. Allow non-enzyme reagents to thaw in a PCR rack on the bench (BSA and polymerase are enzymes stored in glycerol, and do not need to thaw before use). Sample reagents can thaw in the 4°C refrigerator or in a covered rack outside the hood.
  • Mix and spin all reagents and keep in the covered "reagents" cold rack.
  • Prepare your master mix in a 1.7 mL tube placed inside the "mix" cold rack. Transfer your aliquot of water to this rack, because it will be needed for your negative control.
  • When the reagents are aliquoted to your mix (including polymerase), cap the mix and return the reagents to their storage location.
  • Change gloves and then place an appropriate number of PCR tubes/plate in the covered cold tray designated for the PCR hood, and briefly label each row of tubes. You do not need to cap the strips, but do keep them covered. Do not lay cap strips on the bench to label them--handle them carefully and cleanly!
  • Vortex and spin the master mix, then aliquot the desired amount to the wells. It is OK to reuse the tip when aliquoting for standard PCR (but not for metabarcoding), even as you touch the tip to the inside of the tube for proper pipetting. Just check your tip for cavitation and change it once per row "just in case".
  • Cover the rack containing the reaction tubes. Change your gloves. Vortex and spin each of your sample tubes. Double check that sample order in the rack physically matches your lab notebook.
  • Add the sample to the appropriate reaction tube and mix by pipetting. Change tips each time!!
  • Close tubes for each cap-strip when completing each row, or seal the plate when all wells are complete. Make sure the seal is tight to avoid evaporation in the thermocycler.
  • Remove reaction tubes from the cold tray and put them in a transfer container. Move quickly to the post-PCR room, briefly vortex and spin each reaction before placing them in the thermocycler (check to ensure there are no bubbles).
  • Pro-tip #2: When opening tubes and pipetting, be cognizant of where your hands go; avoid manipulating tubes over your master mix or reaction tubes, because this is where most contamination is likely to occur (aerosols or splashes when opening a tube, condensation or ice dripping from the bottom of a tube; dust and microbes falling off your wrists). I usually load my pipette tip with my right hand and use my left hand to open a tube, I carry the pipette from right to left toward the tube but just slightly in front of the cold block where my master mix or reaction tubes are being held so that the source tube never moves far from its rack; to open screw-cap tubes I'll use my right index finger and thumb while holding the pipette ready to go. This strategy may take practice, and it is a GOOD IDEA to practice when no samples or reagents are present.

PCR Troubleshooting

Thermocycler organization

  • The lab maintains a "standard" folder. This folder contains all protocols found on this wiki, only protocols found on this wiki, and is updated along with the protocols on this wiki.
  • Individual users can maintain separate folders with protocols that are used temporarily or for troubleshooting. These protocols must be documented and accessible to other lab users as needed, and will be incorporated to the wiki and standard folder when optimization is complete.

Standard Primer Sets

Below are some useful PCR recipes using reagents that we have (or have had) in the lab for various primer sets and sample types. Please report optimizations and troubleshooting to the lab so that we can make adjustments as needed. These protocols should work, but please do think critically about how they should be adjusted to the needs of your particular project.

Mammal Barcoding

1. Mitochondrial DNA

Plant Barcoding

1. Chloroplast DNA. These protocols mostly use NEB polymerase, simple thermocycling protocols, and sequence well with standard 'pre-mixed' options at GeneWiz. The matK and trnH-psbA protocols could be converted from Phusion for more regular use.

Invertebrate Barcoding

1. Mitochondrial DNA

Illumina Primer Sets

  • Illumina sequencing primers should be engineered with appropriate overhangs for incorporation of sequencing adapters according to URI's standard workflow and the Nextera protocol.
  • Metabarcoding protocols should employ high quality polymerases. This means they should minimally be hot-start polymerases (inactive initially at room temp). We have often used Amplitaq Gold II. Other polymerases that also include proof-reading and high-fidelity features are desirable.


The trnL-P6 marker is one that we have used a lot for dietary DNA analyses. Most of our projects rely on primers g/h but we dabble in using primers c/h. We have used several polymerases for various projects, but it is critical to always maintain consistency within projects. The recipes below correspond to several current/recent projects. Discuss with TRK before starting any new projects, as we are currently moving to a new approach.
Primers g/h:

  • Amplitaq Gold II polymerase: trnL-P6 g/h protocol: Amplitaq. This protocol has been used for Kenya small mammal diet analyses and most of our earlier large mammal diet analyses. Note that this is a 15 uL recipe, which we used for small mammal diets; our standard for large mammal diets ranged from 12.5 uL to 20 uL (see, e.g., Kartzinel et al. 2015; Kartzinel et al. 2019). Note also that this protocol worked really well with the shorter MID-tags that we used in our published work, but did not readily transfer to the longer overhangs required for a Nextera protocol; we could get strong amplification based on primers engineered to have these long overhangs by boosting the Mg concentration to 3.5 mM, but as of mid-2021 we have not generated sequence data based on amplicons generated using this level of Mg.
  • Platinum Taq polymerase: trnL-P6 g/h protocol: Platinum. This protocol has been used for Yellowstone large mammal diet analyses projects on elephants and giraffes in Kenya. We have found it works well with the Nextera overhangs on these primers.

Primers c/h:


  • The IN16STK primers are designed to amplify invertebrate 16S rRNA (DNA sequences) from vertebrate diets. We have both the original IN16STK and the more recent degenerate IN16STK-mod primers available in the lab.
  • [Other experimental COI]


The 16S V4 marker is one that we have used for several microbiome studies. It's critical to always maintain consistency within projects, but we have used several approaches across projects over the years. The recipes below correspond to several current/recent projects. Discuss with TRK before starting any new projects, as we are currently moving to a new approach.
Primers 515f/806r:

  • We have used an Accuprime Mastermix protocol to analyze the gut microbiomes of small mammals in Kenya and large mammals in Yellowstone: 16S-V4 rRNA protocol. These primers are designed to work with the Nextera library prep method.
  • The long Earth Microbiome...

Thermocycling Protocols

  • All thermocycling protocols happen in our post-PCR room, and reactions stay there. No exceptions.
  • All thermocycling protocols include a heated lid (unless otherwise noted).
  • Note that a 4ºC overnight may cause condensation on the thermocycler's block. Please do not allow a hold to really be "Forever"...

Gel Protocols

Many people are handed a gel running protocol without the opportunity to think about how suitable parameters were identified. This can lead to trial and error when developing a new protocol, transferring to new equipment, or joining a new lab. Here are a few useful resources for planning new gel running protocols, identifying common problems, and resolving the issues efficiently.

Below, we list protocols that work for our current gel system. If you have trouble, review the information in the links above and check with the PI.

Storing and reusing materials

  • Gel stain: we only use GelGreen in the lab. This dye is light-sensitive and must be kept in freezer box and wrapped in foil. It can be stored at room temperature. It must be vortexed and spun before each use.
  • TBE buffer: Use the carboy labeled “Used TBE” to fill the gel rig.
  • Agarose gels: Use the carboy labeled “Fresh TBE” to cast your gel.
  • Loading dye: we use NEB Gel Loading Dye Purple (6X). This is a mix of glycerol and a visible stain to allow you to track the progress of your gel run. Stocks and working stocks can be stored at room temperature. When preparing runs with many samples, it is OK to aliquot ~100 μL to each well of a strip tube and use a multichannel pipette to aliquot the appropriate amount to your parafilm.
  • Amplicons are relatively stable at room temperature, so these can be handled at room temperature for the ~15 min it may take you to load a gel. But they should be stored in the post-PCR -20°C freezer between uses.

Pouring a gel

Pore density in gels is an important, and different densities can help visualize DNA strands of different lengths. These densities are determined based on the % of agarose in a gel (usually in the range 0.5-3.0%). For example, to prepare a 1% gel, weigh 0.5 g of agarose and dissolve it in 50 mL of fresh TBE buffer. Because our most commonly used gel rig is small, only 50 mL of agarose needs to be prepared.

  • To visualize PCR results for most metabarcoding sequencing applications in the lab, use a 2% gel (useful for separating ~100-2000 bp amplicons).
  • To visualize PCR results for most Sanger sequencing applications in the lab, use a 1% gel (useful for separating ~400+ bp amplicons).

To prepare a gel:

1. Zero the balance with a weighing paper on top. Weigh the appropriate amount of agarose in grams, and put it into a flask.
2. Measure the appropriate volume of 1x TBE buffer from the “Fresh TBE” carboy into a graduated cylinder.
3. Slowly pour the buffer into the flask containing the agarose, swirling throughout the addition to ensure the agarose gets suspended with minimal clumping.
4. Place the flask in the lab microwave (uncovered) and heat for approximately 2 min. Stop the microwave to swirl the flask using an ovenmit (careful, the flask will be HOT!) every ~30 seconds. Watch the flask carefully because when the agarose is nearly ready it will be close to boiling over. You should stop the microwave and swirl the flask to prevent boiling over when bubbles rise toward the top. When the solution reaches that near-boil-over point, visually check to ensure the solution is fully clear and free of agarose clumps.
5. Place the flask on the counter and quickly add 1:10000 of the GelGreen stain to the solution and swirl the flask 2-3 times to mix thoroughly. If you prepare 50 mL of solution, for example, you should incorporate 5 μL of stain.
6. Place appropriate gel combs in the gel form, thinking carefully about whether you want the large or small wells, and ensure the gel form is on a flat surface (use a bubble level to confirm; don't just eye-ball this).
7. When the agarose cools to ~65 °C, pour the gel into the form. You know it is approaching the right temperature because you can safely lift the flask without an ovenmit, but it becomes uncomfortable to hold it in your hands for more than ~3 seconds at a time. Usually this takes 5-10 min. Pour the gel steadily from a single position near the front-end of the form (do not move the stream around, lest you introduce temperature inconsistencies or bubbles; avoid letting the last drops of solution dribble into the gel form). Large bubbles may be fixed using a clean pipette tip, but small or persistent bubbles may not be worth the trouble to correct.

Loading a gel

  • Once the gel is set, put it in the chamber with the wells facing away from you (near the black electrode: remember, "run to red").
  • Carefully remove the comb and fill the chamber with TBE buffer (this buffer can be used several times -- see section on storing and reusing supplies -- but should be replaced every 2 weeks or more often if conducting many long runs). The buffer should rise to the "fill line" and be <1 cm above the top of the gel.
  • Select a ladder based on the expected size range of the DNA in your samples. Some ladders are not very effective with GelRed/GelGreen. Brian can advise on appropriate ladders for use in the lab, and will include this information here.
  • Cut a strip of parafilm and pipette 1.5 µl of loading dye corresponding to the number of your samples plus one for a DNA ladder. This can be done using a multi-channel pipette if the dye is aliquoted to a strip tube. Pipette slowly as the loading dye contains glycerol and is prone to cavitation, and prone to damaging pipettes through clogging (or getting on your gloves).
  • Add 2 µL of PCR product to the corresponding drop of loading dye, mix (slowly) by pipetting, and then using the same tip you can inject it into the corresponding lane of the gel. This can be done with a multichannel if the dye is applied to the parafilm using a multichannel to ensure the spacing is the same.
  • Preferably, the first and last lane of the gel should be reserved for the ladder.
  • Change tips between samples.

Running a gel

Select voltage based on the side of bands you aim to separate and the dimensions of the gel rig itself (distance between electrodes).

The inter-electrode distance for our gel rig is 19 cm, and we use TBE buffer. 

  • For Sanger and metabarcoding amplicons (<1000 bp), the rule of thumb is 5 V/cm = 19 cm x 5-V/cm = 95. We recommend 120 V for 30 minutes.
  • We use three ladders in the lab, and these should be matched to the expected size of your product. Each ladder comes with a recommended range of V/cm that should also fall between 5-10 V/cm.

Imaging a gel

  • To transfer a gel from the rig to the imager by opening the imager, lifting the gel form out of the buffer, carrying it over to the imager, and carefully sliding the gel onto the glass surface.
  • Position the gel in the viewfinder using the white light and manipulate settings until the image is clear.
  • Turn on the transluminator and manipulate the exposure until you can visualize your ladder and/or bands.
  • Save the image to a USB memory stick, and save them to LabArchives (even if you are just running a "yield gel", we often refer back to these images for troubleshooting).
  • Use a kimwipe (NOT a paper towel) to clean the transluminator surface (can be disposed in regular waste bin).

Disposing of gels and buffers

  • Agarose gels can go in the red lined box in the post PCR lab. They simply desiccate.
  • Spent buffers can go into liquid hazardous waste (empty containers for waste disposal are usually saved next to the red hazmat box in the pre-PCR lab, and these containers can be brought over for use as needed).
  • If a new container is required, fill out the orange chemical hygiene form and affix it to the container. Since the only liquid waste we will produce in this room is TBE buffer, you can label “100 %Tris Borate EDTA buffer”. The Lab Care Manager will contact EHS for disposal when full.

Targeted Sequencing

Targeted sequencing, also called Sanger sequencing, is the protocol we use to sequence single-band/single-product amplicons. We perform sequencing in both directions.

Amplicon clean up

1. ExoSapIT. We use a protocol that dilutes ExoSapIT reagents for cost savings, without typically reducing sequence quality. Diluting the PCR product can often (but not always) also help by reducing saturation of the BigDye signal. Although we may occasionally use the manufacturer's full-strength protocol or hire companies like GeneWiz to prep challenging templates, our default protocol involves an ExoSapIt dilution AND a template dilution. We typically submit these as a "pre-mixed" submission to GeneWiz via Brown for Sanger sequencing.
Run the ExoSapIT cleanup; this creates 7 μL of cleaned amplicon, and we use 3 μL each (6 μL total) for forward and reverse sequencing reactions:

  • Remove ExoSAP-IT reagent from –20°C freezer and keep in cold block throughout procedure.
  • Create an ExoSapIT master mix: for each reaction (N=1), use 5.8 μL water (molecular grade from the post-PCR freezer) and 0.2 μL ExoSapIT enzyme.
  • Aliquot 6.0 μL of this master mix to strip tubes (or plate).
  • Add 1.0 μL of your amplicon corresponding wells of the strip (or plate).
  • Complete incubation in the thermalcycler using the "ExoSapIT-45min" protocol. This protocol includes 37°C for 30 min, followed by 80°C for 15 min, and a 4°C "forever" hold. Use the heated lid.
  • The amplicons are now cleaned up and suitable for a BigDye reaction that can be performed in our lab or by a company like GeneWiz.

Create the diluted "pre-mixed" sample (forward and reverse) for submission; this creates 15 μL for both forward and reverse primers as required by GeneWiz:

  • Thaw or create aliquots of 5 μM "sequencing stocks" of primers and keep them on ice.
  • If you need to create aliquots of sequencing stock primers, make them from your PCR primer stocks: typical working stock primer concentrations for PCR are 10 μM, so this is a 50% dilution (e.g., 50 μL of a 10 μM working stock primer + 50 μL molecular grade water).
  • For each direction you need to sequence (forward and reverse) create SEPARATE tubes with the following ratios:

7 μL molecular grade water; 5 μL sequencing primer (forward OR reverse); 3 μL of the ExoSapIt-cleaned product.

  • Tubes are ready for BigDye reaction, and can be submitted to GeneWiz using their "pre-mixed" protocol and data management system.

Other targeted sequencing steps

Pending decisions to standardize and streamline sequencing protocols for the lab; multiple options may ultimately be available, depending on the urgency and scale of a sequencing project...

  • Chain-terminator (BigDye) reactions:
  • Submission for sequencing.
  • Sequence assembly and analysis.
  • Submitting to BOLD
  • Submitting to GenBank

Illumina MiSeq At URI

Amplicon sequencing

Please coordinate with Tyler at each step
To submit samples for sequencing at University of Rhode Island (URI):
1) Request quote, purchase order, and provide sample information.

  • A) Provide the Orders Contact the following information:
  • -The number of samples to process.
  • -The stage of preparation of the samples (extraction products, PCR products (purified/unpurified), ready-to-load libraries).
  • -A spreadsheet containing a list of sample identifiers corresponding to the labeling of your samples.
  • -The excel file should be formatted with two columns: (i) "Tube Number"; (ii) "Sample Name" (i.e., TK#)
  • B) Once this information is in order, the Orders Contact will work with you to send the quote from URI to the IBES Financial and Grants Coordinator (Paula) to request a purchase order (PO) and send the sample information to the core manager at URI.

2) Prepare samples for shipping.

  • A) URI requires that samples be submitted in PCR strip tubes with individually attached lids. They should be clearly labeled with the same identifiers provided to the lab ordering contact.
  • B) If you require dry ice, coordinate with Brown Environmental Health and Safety (EHS) to ship your samples:
  • -Provide a completed shipping request form (see attached model) submitted 72 hours in advance of the ship date.
  • -Provide a MSDS for carbon dioxide (attached).
  • -Make an appointment to meet with an EHS to package the shipment (they want these scheduled early in the week).

C) Request that the Orders Contact provide a FedEx waybill for your shipment.

  • -This must indicate that the shipment has dry ice and the amount.

D) Meet with EHS to package your shipment having prepared the following in advance:

  • -A cooler and cardboard box with no labels on it (peel them off boxes you want to re-use).
  • -Packing tape.
  • -Your samples.
  • -A copy of your shipment request form, MSDS for dry ice, and FedEx waybill.