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 read through the wiki before beginning lab work or when shifting to a new phase of your lab work if it's been a little while, because policies and protocols are subject to change as we continually improve our work. If details are missing, do not assume they are unwanted; consult with Tyler to identify appropriate pilot protocols and remember to report successes for inclusion on the wiki.
- 1 Pre- and Post-PCR
- 2 General Guidelines
- 2.1 Cleaning lab resources
- 2.2 Equipment Use & Maintenance
- 2.3 Reagent storage and handling
- 3 Waste Disposal
- 4 DNA sampling & extraction protocols
- 5 PCR protocols
- 6 Gel Protocols
- 7 Targeted Sequencing
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 vehicles. These are racks, coolers, and containers 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 vehicles 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. This is where the transfer vehicles will reside, and they must be bleached regularly.
- The transfer station in the post-PCR room is designated on the metal rack near the door.
Cleaning lab resources
- All lab users are responsible for the cleaning and maintenance of each piece of lab equipment each time they use it, including switching off (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 overnight, please do not forget to return and finish the clean up the next day.
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.
- 10% Bleach: Make a fresh solution daily in the bleach-specific squirt bottles. Pour about 10 mL bleach and fill to 100 mL (i.e., adding 90 mL) with tap water (measure with beakers/graduated cylinders).
- 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.
- To wash pre-PCR tube racks and trays, fill a wash basin with 10% bleach (made fresh daily). Make sure items are fully submerged (no air bubbles) and that anything sticky is scrubbed off. After at least 30 min (i.e., end of day) rinse with tap water, shake out, and allow to dry on the racks. When dry (e.g., following morning) 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.
Equipment Use & Maintenance
It is each lab member's responsibility to read equipment manuals.
Pipettes are vital to the quality of your work, and they are expensive. Practice good technique at all times. If you have not used a pipette before (or even if you have used them extensively in other labs), please adopt our general guidelines.
- Avoid dropping pipettes. Place them gently in their holders when not in use.
- Place pipettes back in their holders rather than on the workbench, even when you are just pausing while setting up reactions, to keep them clean and out of danger of spills/contamination.
- Do not turn pipettes past their limits (crank them slowly when approaching limits).
- 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).
- Use only filter tips for pipettes (only filter tips should be ordered for the lab).
- 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!
- Use requires training from a senior member of the lab.
- 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.
- This cabinet should only be used for sample manipulation and DNA extraction. No PCR reagents or PCR products should ever 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, and then finally turn off the hood.
- Always clean and stow the cabinet after every 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).
- 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. The PI or postdoc (others only with training and express permission) will provide project-specific aliquots of these reagents for each experiment.
- 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.
- Stock bags of tubes, strips, and plates can be found in the cabinet over the main bench and behind the hood. The PI or postdoc (others only with express permission) can provide project-specific subsets based on the minimum number anticipated for an experiment (more can be obtained if more are needed, but we must minimize waste).
- 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 floodlights. Tube racks and bags intended for use can be added at this time to be bathed in UV.
- 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 floodlights 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.
- 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)
Bench side freezer
- There is a -20°C "mini" freezer 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.
- If you will go more than ~1 week without using a particular set of samples or reagents for PCR setup, please return them to appropriate long-term storage locations.
- Lab stocks should not be kept in this freezer. Long-term 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.
- (One day we may also have a 4°C refrigerator at the PCR station)
- 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.
- How to select which assay kit to use for samples, PCR products, etc... (Comments, BRPB?)
- 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.
- 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.
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.
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 two 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.
- 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 these 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. 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 sensitive reagents that are not enzymes include Qubit standards, dNTPs, ...
- Common light-sensitive reagents include Qubit reagents, gel dyes...
- Light-sensitive reagents should be left in their storage box until use, and returned promptly.
- Before resuspending primers, clean the PCR cabinet and obtain a fresh aliquot of molecular grade water.
- 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 mass 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.)
- Note that "Lab Ready" primers do not require this preparation before use, as we receive them at 100 μM concentration.
- 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 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 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.
DNA sampling & extraction protocols
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.
Xpedition Buffer vs. Standard Buffer
[Details on why we choose one vs. the other; ordering numbers/information for colleagues to reference] ...
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: (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).
Qiagen Plant Mini Kits
A PCR contains the following necessary reagents:
- PCR-buffer. Salt and pH-stabiliser. User stock of 10x is 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 of 25mM is 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 2.5mM of each dNTP) is kept in your personal PCR box.
- Primers. Single stranded DNA (oligonucleotides), usually 18-30 bp. Stock solutions are kept at 100µM, 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 10-100µg/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).
- 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).
- 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 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.
Thermocycler organization and rules
- 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 based on reagents that we have (or have had) in the lab. They can be matched to primer sets and sample types that we have used with them successfully (below). Please report optimizations and troubleshooting to the lab so that we can make adjustments as needed. Please feel confident that these protocols should work, but please don't hesitate to think critically about how they should be adjusted to the needs of your particular project.
1. Mitochondrial Control Region/D-Loop (Ntie et al., 2010)
Useful for identifying mammalian herbivores based on fecal DNA. Target size = 600-750 bp.
Primers (modifications from Hoelzel et al. 1991 and Shields & Kocher 1991):
N777 - TACACTGGTCTTGTAAACC
H16498 - CCTGAAGTAGGAACCAGATG
PCR Recipe: Amplitaq Gold II 20 μL.
Termocycler Program: Standard 48.
1. Chloroplast trnL (Tablet et al. 1991)
Spans the trnL-P6 loop that is so widely used for DNA metabarcoding. Target size +/- 600 bp.
trnL(UAA)c - CGAAATCGGTAGACGCTACG
trnL(UAA)d - GGGGATAGAGGGACTTGAAC
PCR Recipe: NEB Taq 12.5 μL.
Termocycler Program: Standard 50.
1. COI Folmer primers (Folmer et al. 1994)
Amplifies various phyla; mostly invertebrate but also some vertebrates. Target size +/- 654 bp. Primers LCO1490 – GGTCAACAAATCATAAAGATATTGG
HCO2198 – TAAACTTCAGGGTGACCAAAAAATCA
Illumina Primer Sets
NEB 12.5 μL
|Reagents (final concentration)||1x reaction (μL)||x-fold reaction (μL)|
|10X NEB Taq buffer without Mg||1.25|
|MgCl2 - 25 mM (2.5 mM final)||1.25|
|dNTPs (200 μM each)||1.00|
|Primer 1 - 10 μM (0.2 μM final)||0.25|
|Primer 2 - 10 μM (0.2 μM final)||0.25|
|BSA - mg/mL (0.1 mg/mL final)||0.10|
|DMSO (4% final)||0.05|
|Molecular grade water||6.28|
|DNA template||2.00||Add AFTER aliquoting 10.5 μL to reaction wells|
Amplitaq Gold II 20 uL
|Reagents (final concentration)||1x reaction (μL)||x-fold reaction (μL)|
|10X amplitaq gold II buffer without Mg||2.00|
|MgCl2 - 25 mM (2.5 mM final)||2.00|
|dNTPs (200 μM each)||1.60|
|Primer 1 - 10 μM (0.2 μM final)||0.40|
|Primer 2 - 10 μM (0.2 μM final)||0.40|
|BSA - mg/mL (0.1 mg/mL final)||0.17|
|DMSO (4% final)||0.08|
|Molecular grade water||11.25|
|Amplitaq Gold II||0.10|
|DNA template||2.00||Add AFTER aliquoting 18 μL to reaction wells|
- 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"...
1. Standard 48
- Note that "48" denotes annealing temperature. Our "standard" protocols follow this general cycling regime, but differ in annealing temperature.
|1 cycle||95ºC||10 min|
|35 cycles||94ºC||30 sec|
|1 cycle||72ºC||10 min|
2. Standard 50
- Note that "50" denotes annealing temperature. Our "standard" protocols follow this general cycling regime, but differ in annealing temperature.
|1 cycle||95ºC||10 min|
|35 cycles||94ºC||30 sec|
|1 cycle||72ºC||10 min|
3. Standard 55
- Note that "55" denotes annealing temperature. Our "standard" protocols follow this general cycling regime, but differ in annealing temperature.
|1 cycle||95ºC||10 min|
|35 cycles||94ºC||30 sec|
|1 cycle||72ºC||10 min|
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 getting good looking gels, review the information in the links above and ask a more experienced lab member (including the PI) to help evaluate your technique (just like peer-review).
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 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.
Targeted sequencing, also called Sanger sequencing, is the protocol we use to sequence single-band/single-product amplicons. We perform sequencing in both directions. (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...)
- Clean up (ExoSapIt):
- Chain-terminator (BigDye) reactions:
- Submission for sequencing.
- Sequence assembly and analysis.