Modified 5/18/2016, JJM

This is a protocol for a “generic” PCR run using GoTaq Master Mix in a final volume of 25 uL. At the end I give specifics for running a 16S PCR reaction.

Step 0: Prepare Template DNA.

This step will differ depending on what you’re trying to amplify from.  In some cases you will work from purified plasmid or genomic DNA.  In general you don’t want these stocks to be more than ~100 ng/uL in concentration.  Often, it is sufficient to start from a raw culture if you’re working with bacteria or yeast.  Most of the time you can use 1 uL of an overnight LB culture of E. coli as template DNA.  You can also work from a colony by first resuspending a good amount of colony material in ~ 50 uL of sterile saline, and using 1 uL of that as your template DNA.  In the latter case, make sure you can see some cloudiness in the saline, otherwise you need to get a thicker blob of colony.

You also need to ALWAYS run negative and positive controls with your PCR.  Your negative control should be milli-Q water.  The positive control should be a template that you KNOW should give you a product.  For 16S PCR we often use a concentrated stock of purified E. coli genomic DNA.  If you see a band in your negative control, you know some element of your procedure contained contamination, and you can’t trust any of your results (re-do the PCR).  If you don’t see a band in your positive control, something was wrong in your procedure somewhere along the way — in this case, you can’t be confident that other samples that don’t have bands are actually negative.

Step 1: Prepare Primers.

Purpose: by mixing both primers in a dilute 10 uM working stock, we simplify the process of preparing our master mix in step 2 below.

  1. The primers that we buy are stored in 100 uM stocks.  They are given numbers and names that are stored in the file “JM Primers” on owncloud.  The primers themselves are stored at -20.  Find the pair you want and thaw them out.
  2. Add 80 uL of autoclaved milli-Q water to an Eppendorf tube.
  3. Add 10 uL of forward primer to the tube. Pipet up and down to mix.  Repeat with 10 uL of reverse primer.
  4. Label this tube with the name and/or numbers of your primers.  Return the original primer stocks to the freezer; keep your working stock in a separate freezer box from the master stock.

Step 2: Prepare Master Mix.

Purpose: by making one larger-volume preparation of Taq polymerase and primer and then distributing it to multiple reaction tubes, the effect of pipetting error is minimized.


  1. Get an ice bucket.  Keep all ingredients cold at all times — do not take Taq polymerase off of ice any longer than you absolutely have to.
  2. Get your primer working stock and a tube of GoTaq Master Mix (it’s green) from the freezer and let thaw on ice.
  3. Meanwhile, in you lab notebook, make a table with columns for “Ingredients”, “Per Reaction”, and “Total”.
  4. Under ingredients, list “GoTaq Master Mix”, “Primers”, and “Water”
  5. Under “Per Reaction”, list “25 uL” for GoTaq, “2.5 uL” for primers, and “21.5 uL” for water.  Notice that this adds up to 49 uL — leaving 1 uL for template DNA.
  6. Under “Total”, multiply each “per reaction” value by the total number of reactions you plan to run PLUS THREE.  For instance, if you plan to amplify 16S from three samples, you’ll right “150 uL” by “GoTaq” — 3+3 = 6, 6*25 uL = 150 uL.
  7. WHAT ARE THE THREE EXTRA REACTIONS FOR?  Negative control, positive control, and one extra one to account for the fact that a small amount of liquid sticks to your pipet tips.
  8. Wipe down your work surface with ethanol and make sure it is very clean. If you are working in a PCR hood place your tips, pipets, and other supplies in the hood and run the UV cycle with the sash down.
  9. If any of your reagent tubes are not completely thawed, it’s okay to warm them in your hands or in your pockets until they are.
  10. Wearing gloves, place the volumes indicated in the right column into a sterile Eppendorf tube in the order Water – Primer – GoTaq.  Mix by vortexing and place on ice.

Step 3: Set up reactions.

Purpose: Here is where you actually put together the PCR reactions you plan to run.


  1. Wipe down your work surface with ethanol. If you are working in a PCR hood place your tips, pipets, and other supplies in the hood and run the UV cycle with the sash down. (IF you did this in the previous step, you don’t have to do it again).
  2. Wearing gloves, get a sterile 0.2 mL PCR tube for each reaction, plus 2 more for controls, and put them in a PCR rack.
  3. Label each tube with a sharpie.  Label positive control “POS” and negative control “NEG”, as “+” and “-” are easy to misinterpret. (they can look like “t” or “1” pretty easily)
  4. Using FILTER tips, put 49 uL of the master mix you made in step 2 into each tube.
  5. Now, put 1 uL of the approrpriate template into each tube.
  6. Seal tubes and place on ice while you get the thermal cycler ready.

Step 4: Run PCR.

  1. Every thermal cycler is programmed somewhat differently.  Ours uses a graphical interface that should be pretty intuitive.
  2. All PCRs start out with an INITIAL MELT step to separate the strands of DNA.  For GoTaq, this is at 95C for 2 minutes.  NOTE: IF YOU ARE AMPLIFYING FROM UNPURIFIED COLONIES OR CULTURES, this step should last 10 minutes — using the high temperatures to lyse the cells, kill interfering proteins, and release the DNA.
  3. The next steps will be repeated 35 times.  First, we do a MELT step at 95C for 30 seconds.
  4. Next, we do an ANNEALING step for 30 seconds.  The temperature is dictated by the primer set and sometimes requires troubleshooting.  If you have no idea, start at 55C and see if that works.
  5. Next, we do an EXTENSION step at 72C, the optimum temperature for Taq polymerase.  In this step, the polymerase extends the primer and synthesizes new DNA.  This step should last for approximately 1 minute for every 1000 bp length of the PCR target.
  6. At this point, you usually put in a “GO TO” step, that says to go back to the MELT step 34 times.
  7. Next, we do a FINAL EXTENSION step at 72C for 5 minutes to finish extending any partial products.
  8. Last, put in a REFRIGERATION step to hold the product at 12C forever, or at least until you can take the products and put them in the freezer.
  9. Once the cycler is programmed, put your tubes in.  If you have less than 6, you need to put some dummy tubes in to keep the lid from crushing your tubes.  Close the lid and turn the cycler on.
  10. When the program is over take your tubes out and either immediately run a gel or else put them in the freeezer until you’re ready to look at them.
  11. Don’t forget to shut the thermal cycler down when you take your products out!  It’s not good to leave it at 12C literally forever.

Step 4: Running a gel

Purpose: To visualize PCR products and tell if they are “clean”

NOTE 1: I often prepare my gel right after I put my PCR reactions in the thermal cycler.

NOTE 2: This recipe is for a 1% agarose 50 mL mini-gel which can be used to observe up to 22 samples (by using 2 rows of 12 wells).  If you have more reactions than this, you can use the larger gel casting trays which take 100 mL or 200 mL of agarose, respectively, so scale up accordingly.  Also, 1% agarose is great for most PCR products, but very large (>5 KB) products should use less (0.7%) agarose, and very small products (<0.5 KB) should use more agarose (2%).

  1. Measure 50 mL of TAE buffer in a graduated cylinder and transfer to a 125 mL Erlenmeyer flask.
  2. Weigh 0.5 g of agarose.  MAKE SURE NOT TO USE THE EXPENSIVE LOW-MELTING POINT AGAROSE unless you need it to do gel extractions.
  3. Heat the flask for 1.5 minutes on “High” in the microwave.  Keep an eye on it to make sure it doesn’t boil over (it usually won’t).
  4. Wearing a heat-resistant glove, hold the flask up to the light and gently swirl the liquid.  If you see any solid material, microwave for another 30 seconds.  Repeat until you have a homogenous, melted solution.
  5. Pipet 5 uL of Gel-Red into the gel and swirl to mix.  Note: this can be replaced with other 10,000X dyes, such as Sybr Safe and ethidium bromide; however if you use the latter you MUST WEAR GLOVES and everything the gel or running buffer touches must be treated as contaminated.
  6. Place a mini-gel tray in the casting tray and tighten it shut.  Pour the molten gel gently into the tray, avoiding forming bubbles.
  7. Put in the appropriate well-forming combs and make sure they are properly seated in the casting tray.
  8. Allow the gel to cool and solidify (usually takes about 30 minutes).
  9. Carefully loosen the clamp on the casting tray and remove the mini-gel. Place it into a mini-gel electrophoresis chamber with the wells on the side closest to the black (negative) electrode.
  10. Fill the chamber with TAE buffer until the gel is JUST covered, withe the level of the buffer maybe 5 mm over the gel.
  11. Pipet 10 uL of DNA marker ladder into one well of the gel.  If you are using two rows of wells, put ladder in one well of each row.
  12. Pipet 5 uL of PCR product directly into separate wells.  Note that this assumes you’re using the GREEN GoTaq master mix, which already contains (green) loading buffer.
  13. Put the lid on the electrophoresis chamber.  Make sure the black electrode on the lid goes on the black electrode on the chamber and vice versa with the red electrodes!
  14. Connect the chamber to the power supply and turn it on. Set it for constant voltage and 80 V.  You can tell if it’s on because bubbles will start rising around the electrodes.
  15. When the dye front has migrated about 3/4 of the way across the gel, turn it off and visualize the gel.  This can be done with the transilluminator in the Wibbels lab or with other means.
  16. INTERPRETING THE GEL:  First, check to make sure you have a band in your positive control and don’t have one in your negative.  Also make sure the positive control band is the right size, by comparison with your DNA marker lane.  Then, check each of your experimental PCR bands.  Are they the expected size?  Are there any extra bands, indicating non-specific amplification?  If so, you might have to perform a gel purification to separate the target band from any “noise”.
  17. GelRed gels can be disposed of in the trash.  Running buffer can be reused (but make sure to keep the lid on the gel chamber so it doesn’t evaporate).

Step 5: Submitting for sequencing.

  1. First, run each PCR product (assuming no “extra bands”) through a PCR cleanup kit. If your band was relatively dim, elute in a smaller volume (at least 25 uL) to concentrate the product.
  2. Determine the concentration of each PCR product.  If the bands were VERY bright, you can do this by spectrophotometry using our plate reader.  Otherwise, you will need to use PicoGreen.
  3. Dilute the sample to the appropriate concentration: 6 ng/uL for <500 bp products; 20 ng/uL for 500-1000 bp products; 30 ng/uL for > 1000 bp products.  If your concentration is too low, run the PCR again with multiple tubes for each reaction, and run them ALL through the same purification tube to concentrate them.
  4. Pipet 20 uL of each sample into an Eppendorf tube.  Seal and label appropriately.
  5. Prepare a 1.6 uM solution separately for both the forward and reverse primers.  Place 49.25 uL of milli-Q water into an Eppendorf tube, and then add 0.75 uL of the 100 uM primary stock of the primer.
  6. Take both your diluted primers and sample DNA to the Kaul center, room 406, to submit.  Your results will be posted here when they are finished (usually less than a week).



The primers we use are 27F and 1522R, which amplify a product about 1500 bp long.  Use 55C for the annealing temperature and 1.5 minutes for the elongation step.  You will need about 30 ng/uL for sequencing, so it’s a good idea to run each reaction in triplicate and purify them all together.