Real-Time PCR in Action

hi my name is lexa skepham i am a molecular biologist with the virology laboratory for the center for veterinary biologics today i'm going to be talking about real-time pcr and doing a demonstration of how to set one up as well as how to do the data analysis first i wanted to introduce you to some of the equipment that we recommend you use first of all this is a biosafety cabinet and we also have in this room a clean hood the biosafety cabinet is designed specifically to protect the user and the work from one another so the air circulates inside the biosafety cabinet pulling air in past the user that goes into this grate that is then circulated through a hepa filter before it comes back down into the workspace and you can see how the the airflow is pulled onto that grate so this is the setup that we generally recommend that you use i'm going to be doing the real-time pcr setup just on a bench top i've been doing pcr in real time pcr for 25 years and i can get away with doing that but we don't recommend that most people do it it will also make filming easier for us so we will move out to my bench where we will film putting together the reaction so i want to show you some of the equipment that we'll be using today first the primary pieces of equipment are the pipettors and we have pipetters that range from 1 mil volumes down to 1 microliter volumes and they use different size tips depending on the volume that you are interested in dispensing we use tips that are dnase and rnase free and that also have a filter in them so that reduces any splashing up onto the pipette itself and protects reactions from getting contaminated that way other pieces of equipment that we will use are 1. 5 mil tubes these are the 1. 5 mil tubes that we do most of our work in they have a hinged lid that snaps shut and is then watertight there are correct and incorrect ways to open these tubes one of the correct ways is with both hands and being careful to not touch the inside of the tube another correct way is if the tube is in a rack you can open it without touching the inside of the tube a third method is with a tube opener which very handily helps you not touch the inside of the tube the way that you should not open one of these tubes is one-handed it's very tempting to hold the tube like this and to open it with your thumb but when you do often enough your thumb is going to hit the underside of the lid and when that happens then you have the dna or whatever was in the tube on your thumb and as you go to close the tube as you touch other things then opening another tube then you have transferred what was on your thumb to that cap and contaminated whatever you're working with next i want to talk about some of the specialized real-time pcr equipment that we're going to use the reactions are set up in 96 well plates that hold about 100 microliters each and it's helpful to have a rack to put them in the plates are labeled a through h and one through twelve after your reaction is in them you can seal them a couple of different ways you can seal them with either optical caps like these or optical tape like this the optical caps will snap down and create a very nice seal the optical tapes you want to make sure when you're adding the optical tape on that you do not touch it with your bare fingers that's true for the optical caps as well so i will generally use that to press it down and then particularly go around the edges to try to get those pressed down um but you can see that sometimes the seal on the edge is not so good so if i have to use an optical tape i often will not use the wells that are on the edge so i'm confined to the wells in the center for that reason i prefer the optical caps we also have a small centrifuge it holds six 1.

5 mil tubes and it's just it doesn't go very fast but it is good for getting droplets from the inside of a tube collected down into the bottom of a tube and so we use this a lot we also have a vortexer which allows you to mix a sample easily after you've mixed it you'll then want to put it in the microfuge and spin the liquid back down we have racks that will specifically hold the 1. 5 mil tubes and we have racks that will specifically hold the micro tighter plates so that's it for the equipment and next we will start getting ready to set up the reactions before you start a reaction it's important to have the paperwork ready to make sure that you have performed all of the steps this is an example of the paperwork that we use um it includes the date and who's done the work what the run actually is so this is the tomlinson equine parvo virus tacman pcr we're using the in vitrogen superscript platinum qpcr supermix udg and that is the master mix that we will be using in this section it describes the setup of the reaction so we have water for a single reaction that would be 2. 6 microliters of water 10 microliters of the mastermix udg 0.

8 microliters of the forward primer and the reverse primer and the probe the primers and probes are all at 10 micromolar and up here we can put in how many samples we want to run so in this case i'm saying 39 samples this worksheet in its electronic format we'll automatically calculate this plus one times this to give this volume of 101. 4 microliters so to set up enough reaction mix for this entire experiment we'll use 101. 4 microliters of water 390 microliters of master mix and 31.

2 microliters of each primer and probe the primers and the probe are described below the sequence and how long the amplicon is to be expected the lot numbers for the primers and probe are also in the table above the thermal cycler that we're going to use its id number is here and when we start the reaction we'll put down what time we started it the conditions for the thermal cycling are in this part of the table initially we will heat the sample to 95 degrees for five minutes and that will disassociate the double stranded dna in stage two there is a 40 there are 45 cycles of 95 degrees for 30 seconds followed by 58 degrees for one minute the optical reading occurs in this step this is the e the annealing and elongation step so as the temperature changes from 95 degrees to 58 degrees the primers will sit down on the single stranded dna the polymerase will then start to extend that single stranded into double-stranded dna for about a minute and the thermocycler will take a picture of the fluorescence during this cycle this part of the page down here is reminiscent of a microtiter plate so a through h 1 through 12 a through h one through twelve and you can see that all of our samples are going to be performed in triplicate so the we have unknowns samples number one through five and they will go into a4 a5 a6 and on down columns one two and three are reserved for our standards we make a standard curve by doing a dilution series of a plasmid with known quantities so we have a plasmid that is 10 to the five copies per reaction a plasmid that is 10 to the four copies three two one and then we will also include a water negative control okay so this is the part where we set up our reactions the first thing to do is to glove up and then to wipe down your work area you're going to use bleach a 10 mixture that is made fresh at least weekly by the end of a week the bleach will have degraded so i tend to make it up every monday morning you want to wipe down your pipettes particularly the barrels and your tip boxes after you've used the 10 bleach you need to do a wipe down with 70 ethanol to remove the bleach from the surfaces okay so the first step for doing a real-time pcr is doing a dilution series on the plasmid positive control the reaction that we're doing today is for equine parvo virus and the plasmid that i'm working with is an equine pervo virus three plasmid that contains the target site on it in a single copy and i have in this tube a concentration of 1 times 10 to the 6 copies of that plasmid per microliter so that means targets per microliter and we can use this in a dilution series then to calculate what our quantification starting quantification is for our unknowns keep everything on ice the diluent that i'm going to be using today is one nanogram per microliter of salmon sperm dna having the extra dna in the reaction with the plasmid helps us to get a nice straight standard curve and while it's tempting to make your dilutions in the smallest possible volume i recommend against that i will generally do 10 microliters of dna into 90 microliters of water or 5 microliters of dna into 45 microliters of water or diluent in this case the salmon sperm dna so i'm going to put 45 microliters of salmon sperm dna into each of the dilution tubes and we're going to dilute from 10 to the five copies per microliter down to 10 to the zero copies per microliter and i will label my tubes as i have dispensed dna into them but not before that so that if i get disturbed partway through my procedure then i know where i left off when i can get back to it okay five microliters of 1 times 10 to the 6 copies of the plasmid into 45 microliters of the diluent and when i put it in i plunge up and down a few times to rinse the tip and i make sure that there's no liquid remaining in the tip afterwards so this will be the 10 to the five dilution and i don't know what the magic amount of time is for vortexing to make sure that you have an even suspension but i tend to vortex for an entire minute when i do this i end up with nice curves it might not be necessary to vortex for an entire minute but i have not run the range of times to test and so this is what i stick with it's a very good idea to spin every tube before you open it when you're doing real-time pcr particularly if the sample has been in the refrigerator or the freezer and may have gotten some condensation on the lid or if you have just vortexed it and you may have gotten some condensation around the lid um opening a tube that has condensation around the lid is a very good way to get cross contamination in your reactions so i make a habit of centrifuging a tube every time before i open it and then you want to balance the microfuge so that your your centrifuge head doesn't get off kilter it's not such a big deal in these but it is a very big deal in the big centrifuges so 10 to the five is ready five microliters of 10 to the five into 45 microliters of 10 to the four again i'm rinsing the tip checking to make sure there's no liquid and vortexing for a minute 10 to the 4. since i vortexed it now i need to microfuse it okay five microliters of 10 to the four into 45 microliters of diluent and a vortex 10 to the 3 5 microliters into 45 microliters of diluent 4 10 to the 2 10 to the 2. five microliters ten to the one okay tenfold dilution series of ten to the five copies of your target gene per microliter down to ten to the zero copies of target gene per microliter a tool here that i use sometimes for vortexing it is meant as a tube floater for when you're boiling samples but i've found that if i turn it upside down if i need to vortex a lot of samples simultaneously i can put the tubes in upside down and then vortex on this point and that's a nice way to not cramp your hand at this point it's a very good idea to turn on your real-time thermal cycler the camera needs time to warm up in it and it's just better to not have your reaction in there while you're doing that [Music] so at this point you can put your dilution series on ice [Music] while you're making your master mix so today we're using the platinum qpcr super mix udg and the udg platform incorporates uracils instead of thymodines into the pcr amplicons if your reaction requires handling of the amplicon's post amplification there's a possibility for contaminating your workspace with that uracil containing nucleic acid the udg master mix includes not just the uracils but also an enzyme that will degrade dna rna that has the uracil in it during the 95 minute denaturation cycle at the very beginning before your your second experiment therefore any potentially contaminating template that was generated from your first reaction cannot cause problems with your second reaction so we just have five components of this master mix the udg water forward primer reverse primer and probe so i will add those components in as calculated by the worksheet we have dna rnas free water that we purchase an aliquot into small tubes to make sure that every time we use water it is it is nucleus free as well as dna and rna free so you shouldn't get any contamination from your water okay the udg super mix [Music] okay one thing that i would like to point out here is that some liquids you'll be pipetting are more viscous than others and act differently by capillary action inside your your pipette tips and so i wanted to show you we have some dye here that has glycerol in it so it should be easy to see when you're sucking up something that has glycerol in it you want to put your pipette tip just past the surface draw it up and as you can see as i'm drawing it up the liquid is wicking down the side of the tube you can also get droplets on the end of your pipette when you dispense the liquid then you're going to get wicking on the outside again so this time i've pushed my thumb down to the first stop but we still have this much liquid in the pipette so then a gentle slow push down to the second stop then gets everything out of the tip the super mix is going to have glycerol in it and so it's going to act like that i'm using different pipettes and different sized pipette tips for the different kinds of volumes so this is a much smaller volume therefore a smaller pipette the forward primer be careful not to touch the inside of the lid our primers we also store aliquoted into small volumes um we have some tubes that are aliquoted at 10x concentration and so you need to check your label to make sure that you're at the right concentration before you dispense and finally the probe the probe we aliquot we we aliquot into 50 microliter volumes we also do a dilution down to the required concentration as our very first step after we've received the primer from the the manufacturer we then dispense into very small tubes which we can then break apart and we only freeze thaw this the one time so we receive it from the manufacturer we add buffer to get it to the correct concentration for our reactions dispense it into small volumes and then freeze it and then when we use it since it's already been freeze thawed once we'll throw it away another thing to know about the probe is that it is light sensitive so when you pull it out of the freezer to thaw it it's a good idea to wrap it in foil why you're letting it thaw also while you're setting up your reaction you want to protect it from light that does not mean that you need to do your reactions in the dark some people think you do and it's very hard to pipet correctly in the dark it just means to not dilly dally while you're setting up your reactions i don't like to vortex my master mix i don't think that vortexing is good for enzymes so i will invert the tube a few times and then give it a spin to get any liquid off of the lid and at this point we're ready to dispense the master mix into the pcr plate our total reaction volume for this reaction is 20 microliters so 15 microliters of that is the master mix and 5 microliters of that will be our template when you're dispensing the master mix it is not critical to get a perfect amount a perfect 15 microliters into each well the camera on the uh on the thermal cycler will take an initial fluorescence reading and subtract all of that out from the background subtract all of that background out the important pipetting is when you add your five microliters of template to your reactions when you're doing that you want to be very very precise and careful so 15 microliters of master mix per well this master mix is going to be a little bit sticky kind of like the super mix was because of the glycerol i will generally get the tip wet by measuring up and down a few times and then i just dispense into the wells to the first stop you might get a little bit of liquid but you don't get a build up and you don't get a lot of wicking so it's not critical at this point to make sure that every tiny little droplet is off of your tip it is however important that if you put your wetted tip into the reaction mix and you see a bubble on the end of the tip just pull the tip out and put it back in and that bubble will probably be gone okay so the master mix is dispensed and the tube can go into the trash at this point i'm going to move to my smallest pipettor it's a p10 meaning that its maximum volume is 10 microliters we're going to be dispensing five microliters of our template into each of these wells okay and i'm going to set my templates out in order i'm right-handed so i generally like to work right to left when i'm loading the plate and as i'm doing that then i will cover over the wells that i have added template to to protect them i will also often because if i'm going to be loading an entire plate i feel like i don't want the uh the probe fluorophore to be exposed to light very much so i can also cover up parts of the plate that i don't want to have exposed to light while i'm working on other parts of the plate for my unknowns i don't know which ones are going to have a strong signal and which ones are going to have no signal so the order in which i add them to the plate doesn't necessarily matter i do know however that my 10 to the 5 10 to the 4 concentrations of my dilution series have high enough copy number that i could easily cross contaminate into my unknowns so at this point i'm going to load my unknowns first and then after those are loaded i'm going to put the lids on one thing that you may also want to do depending on how you're loading your wells which direction you're loading your wells and how you plan to seal your plate if you've filled up a row with template you can either cover it with foil or you can rip off a strip of cellophane tape and just put that over the top so that you feel more comfortable that you haven't cross contaminated your sample okay so five microliters you can see on the tip there are marks where the volumes are and so i'm going to very carefully put the tip just past the surface of the the dna sample before i draw it up and then dispensing and i do at this point after i've dispensed i can see a tiny bit of liquid on the end i want that in the reaction so i will touch the side of the tube of the well with the tip and plunge down to the second stop to get that last bit of liquid out of the tip i also don't use the same tip for replicates you'll see a new tip every time i go into my sample that's for a number of reasons first of all every time you use the tip you increase the chance of accumulating liquid that could be transferred to the next the next sample and skew your copy number readings also if you have touched something with your tip while you're dispensing then if you take that same tip and put it back into your sample then you have contaminated that sample and this is really a precious sample so you don't want to do that so new tip every time you go into your [Music] unknowns [Music] the unknown templates are dispensed into the wells and now i'm going to put caps on and i don't want to get the caps contaminated they're hard to get out of the bag without contaminating without touching them and so i tend to use tweezers to pull them out of the bag and i put the caps on the row closest to me first so that my hands aren't hanging over the reactions while i'm putting the lids on for these optical caps it's okay to touch the lids to push them down if your hands are gloved but if you are not wearing gloves then the oils from your skin can get on that surface and cause problems with the fluorescence readings during thermal cycling so i'm going to move on to adding the templates for our standard curves i'm going to switch my foil so that my unknown samples are covered protected from light and i'm going to start at the lowest concentration in this particular case that is my water no template control all of these i'm performing in triplicate okay five microliters of 10 to the zero copies per microliter means i'm going to be adding roughly five copies of my target per reaction [Music] there is some stochasticity associated with pipetting and so the likelihood that you're going to get five and not four or six is not always a reasonable expectation okay 10 to the 1 5 microliters 50 copies per reaction the more concentrated you get the easier it is to have your data points being close together because the difference between 48 or 49 and 50 is not as great as the difference between four and five [Music] okay okay and now we're ready to put the reactions into the thermocycler when you want to spin down a plate you need to make sure that the the bottoms of the tubes are facing out so in this case the tubes are slanted to the outside as it spins and in this piece of equipment here this is the balance plate the uh the bottoms of the tubes are facing me the top of the plate away so i'm going to place it in the spinner this direction just give it a quick spin [Music] that does a couple of things it collects all the liquid to the bottom but if there happened to be bubbles in the very bottom they would act as insulators and so when you're thermal cycling the plate the liquid that was in a tube that had a bubble at the bottom wouldn't actually get the thermal cycling so the spinning pulls the liquids down forces the bubbles up to the top where they don't create a problem we use a bio rad cfx 96 there are a lot of different kinds of real-time thermal cyclers that you can use this is the one that we have and this is where you set the plate this is a heated lid that moves up and down part of part of that sound was the machine adjusting how far away from the lid that that plate was um all of the fluorescence and camera equipment is in the top and there are there's a camera inside that will take pictures of the plate as it's cycling okay so i'm going to go to saved files because this is a protocol that we use quite a bit so 95 degrees for 5 minutes followed by 30 seconds and 95 degrees one minute at 58 degrees and then it cycles back so 45 cycles of 95 and 58 with a plate read at the end of the 58 degree cycles press run and the reactions are 20 microliters if your volume was different you could change that the lid temperature you always want to have at about 105 degrees so that's perfect if you you have choices here for what colors you want the cameras to detect we have a fam fluorophore on our probe and so we can just choose the cyber fam setting okay and the run is starting okay so the lid is preheating up to 105 degrees once it gets there then the machine will start to heat the plate up to 95 degrees for its initial five-minute denaturation while this is happening you can look at the graph right now there's nothing but as the cycles progress the fluorescence that is detected by the cameras will be recorded here and you can actually see the curves develop over time so here's our raw unprocessed data and we have four different views of the same data so that we can refer to different pieces of the information that we need quickly the first thing that i'm going to do is go into view edit plate this is the plate and change the setup to mirror what we actually did so i'm just going to quickly all of these are known and label them with fam and then these a through f where our standards we have three reps for each so replicate series there are three reps starting at replicate one the reps are horizontal and so apply so now you can see our dilution series in three reps and we're going to label these with the number of copies that we put into each well so go to dilution series our first well had five times ten to the fifth copies of the target plasmid in each well so the plasmid that we used was 1 times 10 to the 5 copies per microliter we put 5 microliters of that into our reaction therefore our starting concentration for these wells is five times ten to the five this number has a dilution factor of ten it's decreasing and so if we apply then 5 times 10 to the 5 4 3 2 1 0 appears and that is correct for our standard control a standard curve g 1 through three was our no template control and then we had five five unknowns that let's see also had replicates so we're applying those and all of these wells were empty so we're going to remove those from the analysis unknown one through five i like to recolor these so the data is easy to see when we go back to look at the curves so i go to trace styles and change the colors in a way that is easy for me to remember apply those changes so standard number one was five times ten to the five copies per reaction and those red curves are all on top of one another and in the standard curve chart on the top right the standards are shown as circles and the unknowns are shown as x's the circles are right on top of one another four five times ten to the five copies as we go down there's going to be a little bit of spread between the reps and that is okay we want that spread to not be greater than .

3 ct so this standard this is five copies per reaction this is a hard one to hit and over on the bottom right the cts for these reps are 37. 3 36. 3 35.

3 we would prefer that those cts be within point three of one another but they are not greater than uh 3. 3 a ct difference of 3. 3 is one log and considering that this is the five copies having a ct one apart from one another is just fine here at 50 copies we did a better job and getting the the reps right on top of each other for our unknowns those reps are also right on top of one another 23.

1 23. 23 so that is quite good the data automatically figures out what your starting copy number is for your unknowns by referring back to the data that you gave it for your standards so if standard two five times ten to the four copies is similar to unknown one standard two the cts are about 22. 6 and the unknowns are calculated according to the knowns so this this uh unknown has 4.

4 times 10 to the 4 copies that it started with the number of copies that it ended with can be seen up here as relative fluorescence units so this graph this is the number of cycles so we ran 45 cycles of the reaction so this is from time zero up until an hour and a half later relative fluorescence units just shows the difference in the amount of light given off by each well and so if it is a perfect doubling of your pcr product then you're not going to get any signal for quite a while but then you're going to get this logarithmic curve and what you're looking for is this inflection point right here where this curve crosses this is the threshold that determines what the ct is the threshold can be dragged up and down and that will change what the cts are so establishing a limit of detection based on ct is dangerous because you can change your ct simply by raising and lowering this threshold what you're looking for though is a pcr efficiency of 100 percent and r squared so the the perfection of your data points for your standards to make this curve you want that as close to 1 as possible but it needs to be greater than or equal to 0. 98 and then a slope of exactly 100 percent pcr efficiency is minus 3. 2 so if we change this threshold we raise it if we lower it the pcr efficiencies change now there is a point in here i'm going to switch the scales from logarithmic to linear and the data are exactly the same but it's just a different way to visualize there it is pcr efficiency of 100 slope of minus 3.

32 that means that the likelihood that you're going to get an accurate quantification for your unknowns is better than if your pcr efficiency is off it's okay for your pcr efficiency between to be between 90 and one hundred and ten percent so you can you can export all of the raw data by going up to the export tab export in excel and i'm just gonna export it to the desktop and you're going to get a set of files most of which do not give data that are useful to you however the quantification plate view results and the quantification summary are both very helpful so quantification plate view results so we told the software that we wanted five times ten to the five starting copies in wells a one through three and so that's what it says four three two one down to five copies per reaction the starting copy number of our unknowns was automatically calculated from where those data points fell on the standard curve related to the standards themselves and so you can tell by looking at the numbers that the unknowns in this case were just a different dilution series of the plasmid that i used for my standard control another data sheet that you get is the quantification summary and this one is nice for looking at things like what your average ct is so it's easy to calculate that number it's also easy to tell whether or not these the ct reps are within point three of one another this is an easy way to export data for a report this software also has an ability to generate report i like to use the notes section here to get the notes to pop up you have to not only click the box but then you have to actually click the word notes also for the notes to appear but i like to have right on the front page information such as the pcr efficiency was 100 percent the r squared is 0. 995 etc and say update report and that information appears in that front page so this is the report that's generated where it's saved what the data is named if you scroll down the protocol the plate set up here are the curves themselves the standard curve including the pcr efficiency the r squared slope and then there is this data table that shows all of the raw data that you saw in the excel file so for each well what the sample was what the ct was the average ct standard deviation so that's nice what the starting quantity was for a standard or the calculated starting quantity for an unknown and also starting quantity means and standard deviations down at the bottom is a chart that shows whether or not you have pcr efficiencies that fall within the required 0.