【分享】一个好的PCR文章（英文）
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A. Pilot error hypothesis.
Background:
This is the most common problem with new people. It frequently happens a couple of weeks after someone is "trained" and when they start to work independently.
Symptoms:
- i) Other people in the lab have the same primers working on the same type of material, using the same reagents, and the same thermocycler.
- ii) You are new at PCR.
- iii) You have no experience with molecular biology.
Common causes and solutions:
- i) Miscalculation of components, especially buffer or enzyme.
- ii) Compare your mastermix receipe with others. Some early published receipes suggest using too much enzyme (say 5 units Taq to a 50 ul PCR reaction).
- iii) Did you remember to include all reagents?
- iv) You did not mix your reagent solutions prior to pipetting (usually leads to gradually worse PCR performance on a day-to-day basis).
- v) Enzyme was added to master mix prior to buffer (you killed the enzyme).
- vi) The master mix wasn`t mixed (common, but unlikely to have a consistent drastic effect; more likely to cause inconsistency).
- vii) Gross miss-measurement usually due to not knowing how to use a pipetter or a defect pipetter. Although, common to novices, it is unlikely to have a consistent drastic effect; more likely to cause inconsistency.
- viii) Oil was loaded on the agarose gel instead of the reaction contents. (If not using heated lid on PCR-block).
- ix) Oil was not added to reactions prior to running PCR. (Ignore if using heated lid on PCR-block).
- x) Wrong PCR program, or you forgot to start the program.
- xi) Wrong day of week. Go home, rest, and try again tomorrow or next week.
B. Template dilution hypotheses.
Background:
Crude DNA extracts are often poorly quantified, degrade over time, and contain inhibitors to PCR. If specific products from multicopy regions are the target (e.g., rRNA genes or spacers) too little DNA is rarely the problem. The more common problem is inhibitors. The simplest solution is usually dilution, but it may not work for RAPDs or single copy regions. Too little DNA could be a problem in the latter cases. RNA can be an inhibitor of some regions (e.g., the 18S gene). In this case dilution may not help much but RNase treatment of the samples will.
Symptoms:
- i) Some extracts work, others do not.
- ii) None of the extracts work with any set of primers.
- iii) Extractions are from a complex matrix (e.g. milk, food, soil, fungal spores etc.).
- iv) Extractions that used to work do not work any more.
Tests and solutions:
- i) For a subset of the samples try a series of dilutions (i.e., 10-, 50-, 200-, 500-fold) from the crude extracts and repeat the PCR reactions with these dilutions. If a particular dilution works best try it on the rest of the samples.
- ii) If the dilution series does not work, quantify the DNA in some of the extracts. This can be done crudely by loading 10 ul of the undiluted extract onto a 1% agarose mini-gel. If DNA is detectable and a multicopy region is the target, lack of DNA is probably not the problem - some type of persistent inhibitor is. If lots of RNA is visible on the gel relative to DNA try RNase treatment of some of the samples, diluting them and rerunning the PCR experiments. If some persistent inhibitor seems to be the problem try cleaning the extracts from the agarose-gel with a DNA-recovery kit.
- iii) If all the above fails try a different extraction protocol. There are many to choose from, see DNA extraction strategies.
- iv) Symptom iv) suggests that the DNA may be degrading over time. If you keep the undiluted miniprep frozen it should keep indefinitely. If the dilution stops working go back to the frozen extract and make a new dilution.
- v) Perform a nested PCR approach.
C. Temperature errors hypothesis
Background:
The most critical temperatures are those for annealing and denaturation. An annealing temperature that is too low results in non-specific amplifications. One too high results in no products or a low yield of product. Denaturation temperatures that are too low (usually <90 deg.C) result in lower yields or non-specific products. If they are too high for extended periods of time they can fry the enzyme and reduce or eliminate the yield. What you program into a machine and what you actually get is usually not exactly the same. Thermocyclers generally sense the temperature of the block in someway; the temperature of the reaction may lag behind the block, and different regions of the block may heat or cool faster than others (especially for short periods). The speed at which the thermocycler switches between steps (ramp speed) may have a significant influence on the PCR performance, in particular for PCR-fingerprinting approaches (e.g., RAPDs, DAF, AFLPs).
Symptoms:
- i) New machine which was never checked independently.
- ii) Old machine, and everyone´s reactions quit working.
- iii) One of the above and repeatability is a problem.
- iv) You just changed the size of your reactions and ran it on the "standard" program.
- v) You are using a different thermocycler than the one used in the original protocol.
- vi) The temperature in the room where you keep your thermocycler is not stable.
Testing and adjustment:
- i) Monitor the temperature in the room where you keep your thermocycler. If it varies more than a few degrees you should consider moving the thermocycler to another room with a more stable temperature regime.
- ii) Use a mock-up reaction, a small wire thermocouple and a chart recorder to monitor temperatures in your thermocycler as it runs through the program you are using. Check different regions of the block to see if all slots are equal. If you find unacceptable temperatures adjust your program until it achieves the desired result.
- iii) Try a different thermocycler.
D. Unique template hypothesis.
Background:
If the template and primer are mismatched, particularly at the 3`-end of the primer, amplification will be reduced or eliminated. This is most common when the organisms are distantly related to those that the primer sequences are based upon. Among more closely related taxa sometimes introns are inserted within the priming site. Introns or large inserts can also be inserted between priming sites making the region too large to amplify efficiently. If the target DNA is present in relatively few copies there may be other sites competing for your primers (competitive PCR), and this may in turn significantly lower the yield. Alternatively, if the templates have a very high CG content they may not denature completely and then will not amplify well. A hot start and nucleotide analogues in the amplification (7-deazo G) are reported to help. A problem that is relatively common is the presence of short repetitive elements within your target DNA. If the target DNA is a multicopy DNA, e.g. organelle DNA, there is a risk of amplifying target DNAs with varying repeat numbers, thus yielding a population of PCR products differing slightly in size because of the variation in repeat number.
Symptoms:
- i) You are using a new primer/template combination and the templates are derived from a group of organisms that are not closely related to others that have been tested with these primers.
- ii) You get a weak and/or fuzzy product that is much larger than the expected product.
- iii) Multiple fragments are amplified.
- iv) Your target DNA is a low copy number DNA (in terms of total amount of DNA in the reaction tube).
Tests and solutions:
- i) If you have little or no product try to lower the annealing temperature - this will overcome some primer mismatch. Try lowering initally 5 deg. increments down to maybe 40 or 45 deg. at the lowest (below that we call it RAPDs). If you have success, but the products are non-specific bring the temperature back up a few degrees.
- ii) If symptom i) is the case try other primers that amplify the same region.
- iii) If symptom ii) occurs there is no easy fix, unless other primers are available that avoid the inserted/variable repeat copy-number region. One solution could be to clone the PCR products and sequence characterize them.
- iv) If non-specific products is the symptom try a higher annealing temperature first. Keep pushing it up until either the extra fragments are eliminated or until amplification is eliminated.
- v) If non-specific products are the problem and the previous step did not work, check your denaturing temperature by measuring it directly (see temperature errors hypothesis). If it is not already in range of 93-95 deg. push it up there, and try again.
- vi) Try a nested PCR approach.
- vii) If non-specific products are the problem and neither of the three previous steps helped there are lots of buffer additives that could be tried: e.g., gene 32 (a single stranded binding protein), glycerol (5-10%), or DMSO. Hot starts would also be worth a try and can be automated with wax pellets.
E. Buffer problems hypothesis.
Background:
The main components of the buffer are very stable and unlikely to go bad, but there are many different buffers used for PCR that vary in significant ways. The most significant components are the [Mg2+] and whatever is used to stabilize the enzyme. In theory [Mg2+] should be optimized for each primer/template combination. In practice this is seldom done, but there is some risk in ignoring it. Higher [Mg2+] generally results in higher yield, but if high enough will often result in amplification of non-specific products. Both EDTA and dNTP will chelate the [Mg2+] and lower its effective concentration in the reaction. Many 10X commercial buffers have [Mg2+] of 10 mM; this is often too low for optimal amplification. The stabilizers are also important. Most prevent the enzyme from clumping into inactive multimers. These are probably not the problem, however, unless you are making your own enzyme, or using a unique buffer/commercial enzyme combination for the first time.
Symptoms:
- i) You make up your own buffer and you just made a new batch.
- ii) You use the buffer supplied with the enzyme and you just switched suppliers.
- iii) You have never used this primer pair with these DNAs before and assumed that the buffer you use for other primer/template combinations will work.
- iv) You just made up new dNTPs or intentionally increased their concentration in the reaction.
- v) Your DNA solutions used in the PCR reactions have a significant concentration of EDTA (say 1mM) in them.
Suggestions:
- i) Check your EDTA levels in the DNA extract added to the reactions. They should never be greater than 0.1mM.
- ii) If you are trying a new set of primers that others have described, compare the published [Mg2+], [Mg2+]/[dNTP] to what you are using. If you are using a brand new primer pair then try bringing up the [Mg2+] in multiple trials in 0.5mM increments.
- iii) Try using the commercial buffer but bring the [Mg2+] up to the levels used in your previous buffer. This approach retains all the mystery stabilizers matched to that particular enzyme prep while bringing the [Mg2+] up to what you have previously determined to work best.
- iv) Experiment endlessly with different concentrations and combinations of various stabilizers: glycerol, DMSO, different pH, acetamide, non-ionic detergents (NP-40, and others). - Many people swear by some of these components. They occasionally help yield or specificity, but are unlikely to resolve a "zero-yield" problem. It is also hard to rigorously test these, because they interact with each other. Therefore, this often degenerates to a trial and error - borderline voodoo approach, but it might be worth exploring.
F. Bad dNTPs hypothesis.
Background:
The nucleotide triphosphates (dNTPs) are probably the least stable component. One should have the 10X dNTPs distributed into small tubes (e.g. 200 ul) that are kept frozen, used a few times each, and then discarded.
Symptoms:
- i) The same tube of dNTPs has been repeatedly thawed and used for weeks.
- ii) A new batch of dNTPs was just made up.
Tests:
Try a new tube of 10X dNTPs in a side by side comparison with your old tube.
G. Bad primers hypothesis.
Background:
When stored in a TE buffer and refrigerated, primers are incredibly stable and unlikely to go bad and be the source of your problems. Even if left out on the bench top for short periods of time (say overnight) they are unlikely to go bad unless nucleases are somehow added to them (via bacterial growth or because you did not wear protective gloves). Primers potentially could be synthesized wrong, or poorly, or more likely, they could be diluted incorrectly after synthesis.
Symptoms:
- i) A new batch of primers was just made up.
- ii) A new dilution of primers was just made up.
- iii) PCR performance has gradually decreased from day to day.
- iv) You do not wear protective gloves when performing PCR work.
- iv) The primer stock got left in your back pocket for a 14 day backpacking trip.
Tests and solutions:
- i) Wear protective gloves.
- ii) Other primer pairs can be used as a control; if they don´t work either it is unlikely to be a primer problem.
- iii) Compare the sequence of the primer you wanted with the sequence on the order form - did you transpose any bases?
- iv) Check the concentration of your primers by taking an O.D. at 260 nm. If it is much lower than expected that is your problem.
- v) If possible check the primer with gas chromatography for a single band - multiple bands indicate incomplete synthesis. You can also check this by labelling the primer and running it into a 8% acrylamide gel. If this is your problem, complain to your oligo supplier, demand a refund and then switch to a more trustworthy supplier.
H. Bad enzyme hypothesis.
Background:
This is the least likely cause. The enzyme is very stable, and most manufacturers probably make a good product these days. Bad enzyme used to be common, but this has probably changed. However, different brands do perform differently, andunit definitions are not always equal.
Symptoms:
- i) You bought brand X for the first time, but see buffer problems hypothesis.
- ii) You got daring and made your own enzyme (not hard).
- iii) You microwaved the Taq tube.
Test:
- i) Check the unit definition of your enzyme supplier.
- ii) Try (an)other enzyme brand(s) in (a) side by side comparison(s).
I. Bad karma hypothesis.
Background:
This is basically the "God is punishing you" hypothesis. It sometimes gains a great deal of favor.
Symptoms:
- i) All rational explanations have been exhausted and yet PCR still is not working for you.
- ii) Persistent feelings of guilt (if you are a Catholic, this symptom could be misleading).
Tests and solutions:
- i) Try bungee jumping. If you survive, God must not be too hacked-off at you.
- ii) Atone for your sins and start over at the top of the flow chart.
- iii) If you end up here next time, have someone watch you next time you set up your PCR reactions.
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The "MORE ABOUT" section:
PCR carry-over contamination = DISASTER!
The first thing you must consider when you set up your PCR laboratory is the potential risk of carry-over contamination of amplified products. PCR is such a powerful technique that even a few molecules of template DNA can be amplified to billions of copies in a single reaction. Thus, amplified products that can be transferred from previous amplifications always represent a potential contaminant to successive amplifications (carry-over contamination). To reduce the risk of carry-over contamination, there are some steps you can take:
Separate DNA extraction, pre-PCR set up and post-PCR examination facilities. (Preferably into three different rooms). Do not move equipment like pipettes, racks, microfuges etc. between facilities.
Always include a negative (no DNA) control in your DNA extraction and PCR set ups.
Each person in the lab should have his/her own set of pipettes and reagents for DNA extraction and for pre-PCR. Reagents should be made up and stored in small aliquots that can be discarded if carry-over contamination is suspected or observed.
Always treat tubes and solutions post-PCR assuming that amplified products are present. Use separate post-PCR pipettes for this work.
Avoid creating aerosols. Aerosols are easily created when pipette tips are ejected, or if pipettes are waved vigorously.
Use filter tips to reduce the risk of transferring DNA between tubes.
If possible, have UV-lamps decontaminate the laboratory whenever people are not present.
If you are performing nested PCR, you need to set up separate pre- and post-PCR environments and routines for this work.
Standard PCR setup scheme:
(We recommend this setup, but compare with your PCR protocol and adjust where suitable).
.....When you set up your PCR reactions there are a few things you can do to simplify your work, and to reduce the risk of introducing errors. If you adjust the concentration of all your reagent stocks (except the Taq polymerase) to a standard 10X concentration (relative to the final concentration in the PCR tube on the thermocycler), you do not have to adjust the pipetter for each reagent to be added. Thus, you reduce the risk of adding the wrong volume to your mastermix. If you add your reagents in a defined order, you also reduce the risk of introducing errors.
.....When you dilute your template DNA you should consider using a standard dilution, e.g. a 200X dilution of the crude DNA. To simplify your PCR setup, the standard concentration should be adjusted so that the volume of template DNA dilution added to your PCR tube is 1/2 of the final volume in the tube on the thermocycler. The other 1/2 of the final volume should then be the mastermix.
.....Now you need to know how much mastermix you should make. Start counting the number of PCR reactions to be performed Q = X + N + P + 1. Here X is the number of samples you wish to amplify, N is a negative control, P is a positive control and in addition the mastermix tube must be included (otherwise you will run out of mastermix before X, N and P has received their amount of mastermix. If the final volume V for each PCR reaction is 50 ul you need to make 1/2 X Q X V ul = 25 X Q ul mastermix, i.e. add 5 X Q ul of each reagent (except Taq polymerase) to the mastermix tube.
.....If you have followed the advice given above, you have everything prepared for a standard PCR setup. Add reagent stocks (with suggested concentrations) in the order in which they are listed below:
- Reaction buffer (10X concentrated from supplier of enzyme), see also buffer problems hypothesis above.
- MgCl2 (25mM if not present in reaction buffer, but see also buffer problems hypothesis above.
- dNTPs (2mM dATP, 2mM dCTP, 2 mM dGTP, 2mM dTTP).
- Primer A (forward primer), 5 uM.
- Primer B (reverse primer), 5 uM.
- Taq DNA polymerase (5 units per ul), approximately 0.5-1 units per 50 ul PCR reaction (1.5 ul per 10 reactions is usually suitable), but this may need some adjustments.
.....Load 25 ul of your template DNA dilutions into the PCR reaction tubes and add 25 ul of the mastermix.
Do you wish to learn more about PCR? Visit: BioGuide - PCR
Competitive PCR:
.....Your primers and DNA polymerase does not know the identity of your target DNA. The primers will bind complementary single stranded DNA wherever they find complementary sequences, and the polymerase will perform elongation reactions wherever they find a primer-complementary-strand complex with a free 3´-OH end on the primer. Thus, if your primers can find a lot of alternative sites to bind, alternatively to your target DNA primer sites, you run the risk of failing to amplify your target DNA due to template competition. In addition, shorter fragments (say 250 bp) usually amplify more efficiently than longer fragments (say 1000 bp).
.....Even if the alternative primer sites are not located so that the elongation reaction yields products that can be amplified exponentially, their presence may exhaust your PCR reaction of primer or nucleotide before your target-DNA has been amplified to a detectable level. If the alternative primer sites are located so that more than one product may be exponentially amplified, there is a chance that you may observe more than one amplified product. But more likely, you will only observe the fragment derived from the template that was most abundant in your template DNA solution.
.....One way to overcome the effect of competition between multiple primer sites is to perform a nested PCR amplification. You can also increase the specificity of your PCR reaction by increasing the annealing temperature (see temperature errors hypothesis), lower the [MgCl2+] (see buffer problems hypothesis), or change primers (see bad primers hypothesis).
Nested PCR:
Nested PCR is a very simple strategy that may be used to improve yield significantly.
.....If your target template-DNA is present in very few copies and/or you have inhibitory substances in your template-DNA solution, you will normally not obtain a distinct single PCR product, unless you have unusually specific primers or you have been lucky. A commonly used strategy to improve the yield in such cases has been to perform two successive PCR amplifications, with the products from the first amplification taking the role as template for the second amplification.
.....If you reamplify using the same primers in both successive PCR reactions, you will frequently obtain fuzzy products (unspecific products). If, however, you in the second reaction use primers binding to internal sites in the target first-reaction-product, you will normally obtain a very distinct PCR product. This concept is called nested PCR.
WARNING !
Nested PCR also means that you are amplifying DNA using a PCR product as a template. You must therefore also take the necessary additional steps to avoid carry-over contamination of PCR products.
WARNING !
Primer design:
When you design primers for PCR amplification there are several things you must take into consideration. The following is an incomplete listing that may help you getting started.
.....First of all, you need to know what part of the genome you wish to amplify, whether the same part of the genome has been sequence characterized in other closely related organisms (search EMBL, or GenBank), and whether your target DNA will be present in single or multiple copies per genome.
.....Your primers should be selected carefully, trying to combine specificity and universality, i.e. you may wish to amplify the same variable spacer region of more than one organism and therefore design primers complementary to conserved flanking sequences but specific only for the group of organisms you are interested in. You must therefore compare available sequence data.
.....Normally, the optimal primers have a length of 15-30 nucleotides (nt), and should yield a product > 150 bp. Short primers may sometimes be very useful, but a shorter primer will find many more perfectly matching complementary sites than a longer primer. By increasing the primer length, you are also allowed to increase the annealing temperature and thus obtain a higher specificity in your PCR reaction. On the other hand, by increasing the primer length without increasing the annealing temperature correspondingly, you allow for more wobble (that is primer-template mismatch). A 50:50 ratio of AT to GC base pairs in the primer-template complex is optimal, but this may not always be obtainable. Primers with more GC than AT base pairs will have a higher melting temperature Tm of the primer-template complex than primers dominated by AT base pairs. Mismatches between primer and template may be tolerated, unless they are located at the primer 3´-end. This end should preferably have a G or a C nt.
.....Your primers should be examined for complementarity because they can form primer-dimers if they have complementary ends. Check also for the possibility of hairpin formation (complementary segments within each primer yields secondary structures that may reduce PCR performance significantly). Compute the approximate Tm , e.g. by using the following formula:
Tm = 2 X AT + 4 X CG
where AT is the sum of A and T nt, and CG is the sum of C and G nt in the primer.
A helpful link to primer design is the PrimerDesign home page
You really should do computer assisted design of primers. Try the Primer3 test release
DNA extraction strategies:
.....The simplest DNA extraction strategy is simply to boil your biological sample in extraction buffer, e.g. by microwaving, followed by a short spin and transfer of the supernatant (the template DNA) to a clean sterile tube.
.....Another simple strategy is to lysate your cells in extraction buffer, spin down cell debris, transfer supernatant to a clean sterile tube and successively add a DNA-binding matrix, e.g. glass particles, magnetic beads or other solid supports. Remove the supernatant, and resusupend the DNA in a suitable buffer.
.....Traditional extraction methods involve the use of SDS or CTAB, high salt, phenol and/or chloroform extraction of proteins, RNase treatment, precipitation of DNA with ethanol and final resuspension of DNA from the precipitate. A wide range of these methods have been published. Their advantage over the previously described strategies is mainly that the DNA you obtain may be purer and yield may also be somewhat higher. However, they often include toxic reagents, are more time consuming, and DNA may be more sheared by repeated mixing and spinning of the samples.
.....All of the above strategies may be combined in various ways.
Background:
This is the most common problem with new people. It frequently happens a couple of weeks after someone is "trained" and when they start to work independently.
Symptoms:
- i) Other people in the lab have the same primers working on the same type of material, using the same reagents, and the same thermocycler.
- ii) You are new at PCR.
- iii) You have no experience with molecular biology.
Common causes and solutions:
- i) Miscalculation of components, especially buffer or enzyme.
- ii) Compare your mastermix receipe with others. Some early published receipes suggest using too much enzyme (say 5 units Taq to a 50 ul PCR reaction).
- iii) Did you remember to include all reagents?
- iv) You did not mix your reagent solutions prior to pipetting (usually leads to gradually worse PCR performance on a day-to-day basis).
- v) Enzyme was added to master mix prior to buffer (you killed the enzyme).
- vi) The master mix wasn`t mixed (common, but unlikely to have a consistent drastic effect; more likely to cause inconsistency).
- vii) Gross miss-measurement usually due to not knowing how to use a pipetter or a defect pipetter. Although, common to novices, it is unlikely to have a consistent drastic effect; more likely to cause inconsistency.
- viii) Oil was loaded on the agarose gel instead of the reaction contents. (If not using heated lid on PCR-block).
- ix) Oil was not added to reactions prior to running PCR. (Ignore if using heated lid on PCR-block).
- x) Wrong PCR program, or you forgot to start the program.
- xi) Wrong day of week. Go home, rest, and try again tomorrow or next week.
B. Template dilution hypotheses.
Background:
Crude DNA extracts are often poorly quantified, degrade over time, and contain inhibitors to PCR. If specific products from multicopy regions are the target (e.g., rRNA genes or spacers) too little DNA is rarely the problem. The more common problem is inhibitors. The simplest solution is usually dilution, but it may not work for RAPDs or single copy regions. Too little DNA could be a problem in the latter cases. RNA can be an inhibitor of some regions (e.g., the 18S gene). In this case dilution may not help much but RNase treatment of the samples will.
Symptoms:
- i) Some extracts work, others do not.
- ii) None of the extracts work with any set of primers.
- iii) Extractions are from a complex matrix (e.g. milk, food, soil, fungal spores etc.).
- iv) Extractions that used to work do not work any more.
Tests and solutions:
- i) For a subset of the samples try a series of dilutions (i.e., 10-, 50-, 200-, 500-fold) from the crude extracts and repeat the PCR reactions with these dilutions. If a particular dilution works best try it on the rest of the samples.
- ii) If the dilution series does not work, quantify the DNA in some of the extracts. This can be done crudely by loading 10 ul of the undiluted extract onto a 1% agarose mini-gel. If DNA is detectable and a multicopy region is the target, lack of DNA is probably not the problem - some type of persistent inhibitor is. If lots of RNA is visible on the gel relative to DNA try RNase treatment of some of the samples, diluting them and rerunning the PCR experiments. If some persistent inhibitor seems to be the problem try cleaning the extracts from the agarose-gel with a DNA-recovery kit.
- iii) If all the above fails try a different extraction protocol. There are many to choose from, see DNA extraction strategies.
- iv) Symptom iv) suggests that the DNA may be degrading over time. If you keep the undiluted miniprep frozen it should keep indefinitely. If the dilution stops working go back to the frozen extract and make a new dilution.
- v) Perform a nested PCR approach.
C. Temperature errors hypothesis
Background:
The most critical temperatures are those for annealing and denaturation. An annealing temperature that is too low results in non-specific amplifications. One too high results in no products or a low yield of product. Denaturation temperatures that are too low (usually <90 deg.C) result in lower yields or non-specific products. If they are too high for extended periods of time they can fry the enzyme and reduce or eliminate the yield. What you program into a machine and what you actually get is usually not exactly the same. Thermocyclers generally sense the temperature of the block in someway; the temperature of the reaction may lag behind the block, and different regions of the block may heat or cool faster than others (especially for short periods). The speed at which the thermocycler switches between steps (ramp speed) may have a significant influence on the PCR performance, in particular for PCR-fingerprinting approaches (e.g., RAPDs, DAF, AFLPs).
Symptoms:
- i) New machine which was never checked independently.
- ii) Old machine, and everyone´s reactions quit working.
- iii) One of the above and repeatability is a problem.
- iv) You just changed the size of your reactions and ran it on the "standard" program.
- v) You are using a different thermocycler than the one used in the original protocol.
- vi) The temperature in the room where you keep your thermocycler is not stable.
Testing and adjustment:
- i) Monitor the temperature in the room where you keep your thermocycler. If it varies more than a few degrees you should consider moving the thermocycler to another room with a more stable temperature regime.
- ii) Use a mock-up reaction, a small wire thermocouple and a chart recorder to monitor temperatures in your thermocycler as it runs through the program you are using. Check different regions of the block to see if all slots are equal. If you find unacceptable temperatures adjust your program until it achieves the desired result.
- iii) Try a different thermocycler.
D. Unique template hypothesis.
Background:
If the template and primer are mismatched, particularly at the 3`-end of the primer, amplification will be reduced or eliminated. This is most common when the organisms are distantly related to those that the primer sequences are based upon. Among more closely related taxa sometimes introns are inserted within the priming site. Introns or large inserts can also be inserted between priming sites making the region too large to amplify efficiently. If the target DNA is present in relatively few copies there may be other sites competing for your primers (competitive PCR), and this may in turn significantly lower the yield. Alternatively, if the templates have a very high CG content they may not denature completely and then will not amplify well. A hot start and nucleotide analogues in the amplification (7-deazo G) are reported to help. A problem that is relatively common is the presence of short repetitive elements within your target DNA. If the target DNA is a multicopy DNA, e.g. organelle DNA, there is a risk of amplifying target DNAs with varying repeat numbers, thus yielding a population of PCR products differing slightly in size because of the variation in repeat number.
Symptoms:
- i) You are using a new primer/template combination and the templates are derived from a group of organisms that are not closely related to others that have been tested with these primers.
- ii) You get a weak and/or fuzzy product that is much larger than the expected product.
- iii) Multiple fragments are amplified.
- iv) Your target DNA is a low copy number DNA (in terms of total amount of DNA in the reaction tube).
Tests and solutions:
- i) If you have little or no product try to lower the annealing temperature - this will overcome some primer mismatch. Try lowering initally 5 deg. increments down to maybe 40 or 45 deg. at the lowest (below that we call it RAPDs). If you have success, but the products are non-specific bring the temperature back up a few degrees.
- ii) If symptom i) is the case try other primers that amplify the same region.
- iii) If symptom ii) occurs there is no easy fix, unless other primers are available that avoid the inserted/variable repeat copy-number region. One solution could be to clone the PCR products and sequence characterize them.
- iv) If non-specific products is the symptom try a higher annealing temperature first. Keep pushing it up until either the extra fragments are eliminated or until amplification is eliminated.
- v) If non-specific products are the problem and the previous step did not work, check your denaturing temperature by measuring it directly (see temperature errors hypothesis). If it is not already in range of 93-95 deg. push it up there, and try again.
- vi) Try a nested PCR approach.
- vii) If non-specific products are the problem and neither of the three previous steps helped there are lots of buffer additives that could be tried: e.g., gene 32 (a single stranded binding protein), glycerol (5-10%), or DMSO. Hot starts would also be worth a try and can be automated with wax pellets.
E. Buffer problems hypothesis.
Background:
The main components of the buffer are very stable and unlikely to go bad, but there are many different buffers used for PCR that vary in significant ways. The most significant components are the [Mg2+] and whatever is used to stabilize the enzyme. In theory [Mg2+] should be optimized for each primer/template combination. In practice this is seldom done, but there is some risk in ignoring it. Higher [Mg2+] generally results in higher yield, but if high enough will often result in amplification of non-specific products. Both EDTA and dNTP will chelate the [Mg2+] and lower its effective concentration in the reaction. Many 10X commercial buffers have [Mg2+] of 10 mM; this is often too low for optimal amplification. The stabilizers are also important. Most prevent the enzyme from clumping into inactive multimers. These are probably not the problem, however, unless you are making your own enzyme, or using a unique buffer/commercial enzyme combination for the first time.
Symptoms:
- i) You make up your own buffer and you just made a new batch.
- ii) You use the buffer supplied with the enzyme and you just switched suppliers.
- iii) You have never used this primer pair with these DNAs before and assumed that the buffer you use for other primer/template combinations will work.
- iv) You just made up new dNTPs or intentionally increased their concentration in the reaction.
- v) Your DNA solutions used in the PCR reactions have a significant concentration of EDTA (say 1mM) in them.
Suggestions:
- i) Check your EDTA levels in the DNA extract added to the reactions. They should never be greater than 0.1mM.
- ii) If you are trying a new set of primers that others have described, compare the published [Mg2+], [Mg2+]/[dNTP] to what you are using. If you are using a brand new primer pair then try bringing up the [Mg2+] in multiple trials in 0.5mM increments.
- iii) Try using the commercial buffer but bring the [Mg2+] up to the levels used in your previous buffer. This approach retains all the mystery stabilizers matched to that particular enzyme prep while bringing the [Mg2+] up to what you have previously determined to work best.
- iv) Experiment endlessly with different concentrations and combinations of various stabilizers: glycerol, DMSO, different pH, acetamide, non-ionic detergents (NP-40, and others). - Many people swear by some of these components. They occasionally help yield or specificity, but are unlikely to resolve a "zero-yield" problem. It is also hard to rigorously test these, because they interact with each other. Therefore, this often degenerates to a trial and error - borderline voodoo approach, but it might be worth exploring.
F. Bad dNTPs hypothesis.
Background:
The nucleotide triphosphates (dNTPs) are probably the least stable component. One should have the 10X dNTPs distributed into small tubes (e.g. 200 ul) that are kept frozen, used a few times each, and then discarded.
Symptoms:
- i) The same tube of dNTPs has been repeatedly thawed and used for weeks.
- ii) A new batch of dNTPs was just made up.
Tests:
Try a new tube of 10X dNTPs in a side by side comparison with your old tube.
G. Bad primers hypothesis.
Background:
When stored in a TE buffer and refrigerated, primers are incredibly stable and unlikely to go bad and be the source of your problems. Even if left out on the bench top for short periods of time (say overnight) they are unlikely to go bad unless nucleases are somehow added to them (via bacterial growth or because you did not wear protective gloves). Primers potentially could be synthesized wrong, or poorly, or more likely, they could be diluted incorrectly after synthesis.
Symptoms:
- i) A new batch of primers was just made up.
- ii) A new dilution of primers was just made up.
- iii) PCR performance has gradually decreased from day to day.
- iv) You do not wear protective gloves when performing PCR work.
- iv) The primer stock got left in your back pocket for a 14 day backpacking trip.
Tests and solutions:
- i) Wear protective gloves.
- ii) Other primer pairs can be used as a control; if they don´t work either it is unlikely to be a primer problem.
- iii) Compare the sequence of the primer you wanted with the sequence on the order form - did you transpose any bases?
- iv) Check the concentration of your primers by taking an O.D. at 260 nm. If it is much lower than expected that is your problem.
- v) If possible check the primer with gas chromatography for a single band - multiple bands indicate incomplete synthesis. You can also check this by labelling the primer and running it into a 8% acrylamide gel. If this is your problem, complain to your oligo supplier, demand a refund and then switch to a more trustworthy supplier.
H. Bad enzyme hypothesis.
Background:
This is the least likely cause. The enzyme is very stable, and most manufacturers probably make a good product these days. Bad enzyme used to be common, but this has probably changed. However, different brands do perform differently, andunit definitions are not always equal.
Symptoms:
- i) You bought brand X for the first time, but see buffer problems hypothesis.
- ii) You got daring and made your own enzyme (not hard).
- iii) You microwaved the Taq tube.
Test:
- i) Check the unit definition of your enzyme supplier.
- ii) Try (an)other enzyme brand(s) in (a) side by side comparison(s).
I. Bad karma hypothesis.
Background:
This is basically the "God is punishing you" hypothesis. It sometimes gains a great deal of favor.
Symptoms:
- i) All rational explanations have been exhausted and yet PCR still is not working for you.
- ii) Persistent feelings of guilt (if you are a Catholic, this symptom could be misleading).
Tests and solutions:
- i) Try bungee jumping. If you survive, God must not be too hacked-off at you.
- ii) Atone for your sins and start over at the top of the flow chart.
- iii) If you end up here next time, have someone watch you next time you set up your PCR reactions.
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The "MORE ABOUT" section:
PCR carry-over contamination = DISASTER!
The first thing you must consider when you set up your PCR laboratory is the potential risk of carry-over contamination of amplified products. PCR is such a powerful technique that even a few molecules of template DNA can be amplified to billions of copies in a single reaction. Thus, amplified products that can be transferred from previous amplifications always represent a potential contaminant to successive amplifications (carry-over contamination). To reduce the risk of carry-over contamination, there are some steps you can take:
Separate DNA extraction, pre-PCR set up and post-PCR examination facilities. (Preferably into three different rooms). Do not move equipment like pipettes, racks, microfuges etc. between facilities.
Always include a negative (no DNA) control in your DNA extraction and PCR set ups.
Each person in the lab should have his/her own set of pipettes and reagents for DNA extraction and for pre-PCR. Reagents should be made up and stored in small aliquots that can be discarded if carry-over contamination is suspected or observed.
Always treat tubes and solutions post-PCR assuming that amplified products are present. Use separate post-PCR pipettes for this work.
Avoid creating aerosols. Aerosols are easily created when pipette tips are ejected, or if pipettes are waved vigorously.
Use filter tips to reduce the risk of transferring DNA between tubes.
If possible, have UV-lamps decontaminate the laboratory whenever people are not present.
If you are performing nested PCR, you need to set up separate pre- and post-PCR environments and routines for this work.
Standard PCR setup scheme:
(We recommend this setup, but compare with your PCR protocol and adjust where suitable).
.....When you set up your PCR reactions there are a few things you can do to simplify your work, and to reduce the risk of introducing errors. If you adjust the concentration of all your reagent stocks (except the Taq polymerase) to a standard 10X concentration (relative to the final concentration in the PCR tube on the thermocycler), you do not have to adjust the pipetter for each reagent to be added. Thus, you reduce the risk of adding the wrong volume to your mastermix. If you add your reagents in a defined order, you also reduce the risk of introducing errors.
.....When you dilute your template DNA you should consider using a standard dilution, e.g. a 200X dilution of the crude DNA. To simplify your PCR setup, the standard concentration should be adjusted so that the volume of template DNA dilution added to your PCR tube is 1/2 of the final volume in the tube on the thermocycler. The other 1/2 of the final volume should then be the mastermix.
.....Now you need to know how much mastermix you should make. Start counting the number of PCR reactions to be performed Q = X + N + P + 1. Here X is the number of samples you wish to amplify, N is a negative control, P is a positive control and in addition the mastermix tube must be included (otherwise you will run out of mastermix before X, N and P has received their amount of mastermix. If the final volume V for each PCR reaction is 50 ul you need to make 1/2 X Q X V ul = 25 X Q ul mastermix, i.e. add 5 X Q ul of each reagent (except Taq polymerase) to the mastermix tube.
.....If you have followed the advice given above, you have everything prepared for a standard PCR setup. Add reagent stocks (with suggested concentrations) in the order in which they are listed below:
- Reaction buffer (10X concentrated from supplier of enzyme), see also buffer problems hypothesis above.
- MgCl2 (25mM if not present in reaction buffer, but see also buffer problems hypothesis above.
- dNTPs (2mM dATP, 2mM dCTP, 2 mM dGTP, 2mM dTTP).
- Primer A (forward primer), 5 uM.
- Primer B (reverse primer), 5 uM.
- Taq DNA polymerase (5 units per ul), approximately 0.5-1 units per 50 ul PCR reaction (1.5 ul per 10 reactions is usually suitable), but this may need some adjustments.
.....Load 25 ul of your template DNA dilutions into the PCR reaction tubes and add 25 ul of the mastermix.
Do you wish to learn more about PCR? Visit: BioGuide - PCR
Competitive PCR:
.....Your primers and DNA polymerase does not know the identity of your target DNA. The primers will bind complementary single stranded DNA wherever they find complementary sequences, and the polymerase will perform elongation reactions wherever they find a primer-complementary-strand complex with a free 3´-OH end on the primer. Thus, if your primers can find a lot of alternative sites to bind, alternatively to your target DNA primer sites, you run the risk of failing to amplify your target DNA due to template competition. In addition, shorter fragments (say 250 bp) usually amplify more efficiently than longer fragments (say 1000 bp).
.....Even if the alternative primer sites are not located so that the elongation reaction yields products that can be amplified exponentially, their presence may exhaust your PCR reaction of primer or nucleotide before your target-DNA has been amplified to a detectable level. If the alternative primer sites are located so that more than one product may be exponentially amplified, there is a chance that you may observe more than one amplified product. But more likely, you will only observe the fragment derived from the template that was most abundant in your template DNA solution.
.....One way to overcome the effect of competition between multiple primer sites is to perform a nested PCR amplification. You can also increase the specificity of your PCR reaction by increasing the annealing temperature (see temperature errors hypothesis), lower the [MgCl2+] (see buffer problems hypothesis), or change primers (see bad primers hypothesis).
Nested PCR:
Nested PCR is a very simple strategy that may be used to improve yield significantly.
.....If your target template-DNA is present in very few copies and/or you have inhibitory substances in your template-DNA solution, you will normally not obtain a distinct single PCR product, unless you have unusually specific primers or you have been lucky. A commonly used strategy to improve the yield in such cases has been to perform two successive PCR amplifications, with the products from the first amplification taking the role as template for the second amplification.
.....If you reamplify using the same primers in both successive PCR reactions, you will frequently obtain fuzzy products (unspecific products). If, however, you in the second reaction use primers binding to internal sites in the target first-reaction-product, you will normally obtain a very distinct PCR product. This concept is called nested PCR.
WARNING !
Nested PCR also means that you are amplifying DNA using a PCR product as a template. You must therefore also take the necessary additional steps to avoid carry-over contamination of PCR products.
WARNING !
Primer design:
When you design primers for PCR amplification there are several things you must take into consideration. The following is an incomplete listing that may help you getting started.
.....First of all, you need to know what part of the genome you wish to amplify, whether the same part of the genome has been sequence characterized in other closely related organisms (search EMBL, or GenBank), and whether your target DNA will be present in single or multiple copies per genome.
.....Your primers should be selected carefully, trying to combine specificity and universality, i.e. you may wish to amplify the same variable spacer region of more than one organism and therefore design primers complementary to conserved flanking sequences but specific only for the group of organisms you are interested in. You must therefore compare available sequence data.
.....Normally, the optimal primers have a length of 15-30 nucleotides (nt), and should yield a product > 150 bp. Short primers may sometimes be very useful, but a shorter primer will find many more perfectly matching complementary sites than a longer primer. By increasing the primer length, you are also allowed to increase the annealing temperature and thus obtain a higher specificity in your PCR reaction. On the other hand, by increasing the primer length without increasing the annealing temperature correspondingly, you allow for more wobble (that is primer-template mismatch). A 50:50 ratio of AT to GC base pairs in the primer-template complex is optimal, but this may not always be obtainable. Primers with more GC than AT base pairs will have a higher melting temperature Tm of the primer-template complex than primers dominated by AT base pairs. Mismatches between primer and template may be tolerated, unless they are located at the primer 3´-end. This end should preferably have a G or a C nt.
.....Your primers should be examined for complementarity because they can form primer-dimers if they have complementary ends. Check also for the possibility of hairpin formation (complementary segments within each primer yields secondary structures that may reduce PCR performance significantly). Compute the approximate Tm , e.g. by using the following formula:
Tm = 2 X AT + 4 X CG
where AT is the sum of A and T nt, and CG is the sum of C and G nt in the primer.
A helpful link to primer design is the PrimerDesign home page
You really should do computer assisted design of primers. Try the Primer3 test release
DNA extraction strategies:
.....The simplest DNA extraction strategy is simply to boil your biological sample in extraction buffer, e.g. by microwaving, followed by a short spin and transfer of the supernatant (the template DNA) to a clean sterile tube.
.....Another simple strategy is to lysate your cells in extraction buffer, spin down cell debris, transfer supernatant to a clean sterile tube and successively add a DNA-binding matrix, e.g. glass particles, magnetic beads or other solid supports. Remove the supernatant, and resusupend the DNA in a suitable buffer.
.....Traditional extraction methods involve the use of SDS or CTAB, high salt, phenol and/or chloroform extraction of proteins, RNase treatment, precipitation of DNA with ethanol and final resuspension of DNA from the precipitate. A wide range of these methods have been published. Their advantage over the previously described strategies is mainly that the DNA you obtain may be purer and yield may also be somewhat higher. However, they often include toxic reagents, are more time consuming, and DNA may be more sheared by repeated mixing and spinning of the samples.
.....All of the above strategies may be combined in various ways.
