Monday, November 21, 2005

A DGGE Guide

DON’T PANIC

A GUIDE TO DENATURING GRADIENT GEL ELECTROPHORESIS

By Stefan J. Green (http://www.stefangreen.com/)
With help from members of the Yahoo DGGE Group
And many other DGGE Users

Version 2 – Updated Nov 28 2005

-- CLICK HERE FOR MOST RECENT UPDATES --

TABLE OF CONTENTS

INTRODUCTION
THEORY
EQUIPMENT REQUIRED
GRADIENT FORMER
PERISTALTIC PUMPS
POWER SUPPLY
PREPARATION OF GELS
Web Protocols (For Bio-Rad)
PCR / NESTED PCR FOR DGGE
DGGE standards
PCR product cleanup prior to DGGE analyses
How much PCR product to load on DGGE?
How much DNA to put in each PCR reaction?
Primer dimerization
Nested PCR
ELECTROPHORESIS
RUN TIME / VOLTAGE
INITIAL VOLTAGE VALUES
AMPERES VS VOLTS
GEL STAINING AND VISUALIZATION
SILVER STAINING
GEL TRANSFER FROM BUFFER TO UV TABLE
ADD GLYCEROL TO INCREASE GEL FLEXIBILITY
GEL ANALYSIS
GEL ANALYSIS PRODUCTS
BAND EXCISION AND ANALYSIS
Avoiding Band Excision by Direct PCR product Sequencing
High-throughput sequencing techniques
HOW TO EXCISE DGGE BANDS
WHAT TO DO WITH YOUR EXCISED BAND
Bead-beating
Gel electrophoresis and cleanup
WHAT TO DO IF YOUR EXCISED BAND ISN’T PURE
A PROBLEMS CHECKLIST
TROUBLESHOOTING AND ADVANCED TOPICS
ACRYLAMIDE PERCENTAGE
BLURRY, FUZZY OR SMEARED BANDS / SUDDEN CHANGE IN QUALITY OF GELS
BUFFER RE-USE
Detection issues / DGGE SENSITIVITY
DGGE ANALYSIS OF CLONES
GC CLAMPS
HIGH DIVERSITY ENVIRONMENTS – MEASURING DIVERSITY
MIGRATION OF BANDS INTO THE GEL IS LIMITED OR ABSENT? (NO VISIBLE BANDS IS A FREQUENT COMPLAINT)
MULTIPLE BANDING FROM SINGLE POPULATIONS AND MULTIPLE SEQUENCES FROM SINGLE POPULATIONS
NON-RIBOSOMAL RNA GENE ANALYSES FOR BACTERIAL COMMUNITY ANALYSIS
PCR ISSUES (There’s no end to these…)
Chimeras / Heteroduplex formation
Degenerate primers
Double bands
PCR Inhibition due to humics and polysaccharides
PCR Contamination
Primer cleanup
Primer Mistmatches / PCR Bias
Primer Design
Quantification
Optimization
General PCR articles
PERPENDICULAR GELS
POLYMERIZATION
SEPARATION OF BANDS
SINGLE STRANDED DNA
SMILING GELS
STORING GELS AFTER ELECTROPHORESIS
WELLS – WAVY, POOR QUALITY, ETC.
DGGE OF HIGH GC PCR FRAGMENTS
FANCY DGGE TECHNIQUES
RECOMMENDED PCR-DGGE PRIMER SETS AND GRADIENT CONDITIONS


INTRODUCTION

Denaturing gradient gel electrophoresis (DGGE) is a commonly used technique in molecular biology and has become a staple of environmental microbiology for characterization of population structure and dynamics. The method is a powerful one, and can rapidly provide a tangible characterization of community diversity and composition, and shifts in population can be readily demonstrated. DGGE analyses are also used in the medical field for detection of mutations, including single nucleotide polymorphisms (SNPs). These advantages are coupled to a number of limitations and these limitations should be well understood before employing the technique. There is often a steep learning curve. The purpose of this page is to provide a source of information for both the beginning and experienced user of DGGE analysis. This site is not meant to be exhaustive, and references to seminal articles will be provided as well as links to other sites with useful information. I would also like to acknowledge the large contribution made by the members of the Yahoo DGGE group (http://groups.yahoo.com/group/dgge/). In large part, this site is a summary of the information provided by members of that community.

While there are a number of trials and tribulations related to the actual operation of the DGGE analysis, it is important to remember that many of the difficulties with DGGE belong to the stages prior to the DGGE. Since DGGE analyses require a significant amount of DNA for detection, a polymerase chain reaction (PCR) must be performed prior to analysis. Thus, all the troublesome features of sampling, DNA (or RNA) extraction, reverse transcription (if employing RNA extraction), PCR primer design, PCR conditions, and PCR cleanup bear some thought when troubleshooting DGGE problems. Every stage of molecular analysis can impact (often negatively) each stage downstream. However, these are considerations that are endemic to molecular biology, and indeed to experimental science as a whole. It has been my experience, however, that DGGE analyses are exquisitely sensitive to PCR problems and thus special attention should be given to this stage. I recommend the following papers for those interested in using DGGE or other any other technique for analysis of microbial diversity, ecology, population structure and population composition:

Morris, C. E., M. Bardin, O. Berge, P. Frey-Klett, N. Fromin, H. Girardin, M.-H. Guinebretière, P. Lebaron, J. M. Thiéry, and M. Troussellier. 2002. Microbial biodiversity: approaches to experimental design and hypothesis testing in primary scientific literature from 1975 to 1999. Microbiol. Mol. Biol. Rev. 66:592-616
von Wintzingerode F, Gobel UB, Stackebrandt E. 1997. Determination of microbial diversity in environmental samples: pitfalls of PCR-based rRNA analysis. FEMS Microbiol Rev. 21(3):213-29

The use of DGGE as a tool for analysis of microbial communities has grown in sophistication. It is no longer adequate or appropriate to present DGGE analyses by themselves as an indication of community change. Sequence analyses are the current currency of molecular analyses and must accompany any DGGE analysis. However, DGGE profiles can be analyzed by image analysis software and community profiles as a whole can be taken for cluster analysis. There are a number of features of analysis of complex communities by DGGE which may limit the effectiveness of these cluster analyses; nonetheless, if done properly, such analyses can be powerful. For acquiring sequence information from DGGE, much attention has been focused on the limited sequence information recoverable from the relatively short DNA fragments suitable for DGGE (empirical studies suggest that 500-600 bp fragments are the maximum suitable for DGGE; this has been exceeded occasionally). I will present here various means to recover sequence information from DGGE analyses, and will include a technique for recovering sequence information that is longer than the PCR fragments used for DGGE.

Finally, I would be delighted to receive any comments from the public at large regarding this site and I will be happy to update the site with new information. I can be reached at sjg172@gmail.com or by leaving notes on this site.

THEORY

DGGE analyses are employed for the separation of double-stranded DNA fragments that are identical in length, but differ in sequence. In practice, this refers to the separation of DNA fragments produced via PCR amplification. The technique exploits (among other factors) the difference in stability of G-C pairing (3 hydrogen bonds per pairing) as opposed to A-T pairing (2 hydrogen bonds). A mixture of DNA fragments of different sequence are electrophoresed in an acrylamide gel containing a gradient of increasing DNA denaturants. In general, DNA fragments richer in GC will be more stable and remain double-stranded until reaching higher denaturant concentrations. Double-stranded DNA fragments migrate better in the acrylamide gel, while denatured DNA molecules become effectively larger and slow down or stop in the gel. In this manner, DNA fragments of differing sequence can be separated in an acrylamide gel.

For a full review of theory and of similar methods such as temperature gradient gel electrophoresis (TGGE) or single strand conformational polymorphism (SSCP) the following sites/articles are provided:

· G. Muyzer, E.C. de Waal and A.G. Uitterlinden. 1993. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl. Environ. Microbiol., 59:695-700. Cited at least 800 times!!!
· Muyzer, G. 1999. DGGE/TGGE a method for identifying genes from natural ecosystems. Current Opinion in Microbiology. 2(3): 317-322.
· Muyzer G and Smalla K. 1998. Application of denaturing gradient gel electrophoresis and temperature gradient gel electrophoresis in microbial ecology. Antonie van Leeuvenhoeck, 73:127-141.
· Liu, W. T., Huang, C. L., Hu, J. Y., Song, L. F., Ong, S. L. and Ng, W. J. 2002. Denaturing gradient gel electrophoresis polymorphism for rapid 16S rDNA clone screening and microbial diversity study. Journal of Bioscience and Bioengineering 93, 101-103.
· Miller, K.M., Ming, T.J., Schulze, A.D. and Withler, R.E. 1999. Denaturing gradient gel electrophoresis (DGGE): a rapid and sensitive technique to screen nucleotide sequence variation in populations. Biotechniques. 27(5):1016-8, 1020-2.
· L.A. Knapp. 2005. Denaturing gradient gel electrophoresis and its use in the detection of major histocompatibility complex polymorphism. Tissue Antigens. 65(3):211.
· D. Ercolini. 2004. PCR-DGGE fingerprinting: novel strategies for detection
· of microbes in food. Journal of Microbiological Methods 56 (2004) 297– 314.
· S. G. Fischer and L. S. Lerman. 1983. DNA Fragments Differing by Single Base-Pair Substitutions are Separated in Denaturing Gradient Gels: Correspondence with Melting Theory. PNAS 80:1579-1583.
· Glavac D, Dean M. 1993. Optimization of the single-strand conformation polymorphism (SSCP) technique for detection of point mutations. Hum Mutat. 1993;2(5):404-14.
· Sheffield VC, Beck JS, Kwitek AE, Sandstrom DW, Stone EM. 1993. The sensitivity of single-strand conformation polymorphism analysis for the detection of single base substitutions. Genomics.16(2):325-32.
· Zhongtang Yu and Mark Morrison. 2004. Comparisons of Different Hypervariable Regions of rrs Genes for Use in Fingerprinting of Microbial Communities by PCR-Denaturing Gradient Gel Electrophoresis. Appl. Environ. Microbiol. 70:4800-4806.
· Melt-Madge (Thermal Time Ramp Gel Electrophoresis http://www.ngrl.org.uk/Wessex/meltmadge.htm

Some users may also want to consider terminal restriction fragment length polymorphism (T-RFLP) analyses for community structure analyses. The advantage of such analyses is that they can provide some phylogenetic information based on restriction fragment analysis, and such analyses can also generate some quantitative data that is less readily available with respect to DGGE analyses. In addition, T-RFLP may be suitable for some functional genes and gene fragments sizers for which DGGE analyses are not suitable. T-RFLP may also avoid some problems with degenerate primers that can plague some DGGE analyses. Some manuscripts and websites to consider below:

· Beatriz Díez, Carlos Pedrós-Alió, Terence L. Marsh, and Ramon Massana. 2001. Application of Denaturing Gradient Gel Electrophoresis (DGGE) To Study the Diversity of Marine Picoeukaryotic Assemblages and Comparison of DGGE with Other Molecular Techniques. Applied and Environmental Microbiology, 67(7): 2942-2951.
· Osborn, A. Mark, Moore, Edward R. B. & Timmis, Kenneth N. 2000. An evaluation of terminal-restriction fragment length polymorphism (T-RFLP) analysis for the study of microbial community structure and dynamics. Environmental Microbiology 2 (1), 39-50.
· S.L. Dollhopf, S.A. Hashsham, J.M. Tiedje. 2001. Interpreting 16S rDNA T-RFLP Data: Application of Self-Organizing Maps and Principal Component Analysis to Describe Community Dynamics and Convergence. Microbial Ecology 42(4): 495 – 505.
· Frag Sorter by Fred Michel: http://www.oardc.ohio-state.edu/trflpfragsort/contact.php

EQUIPMENT REQUIRED

I do not explicitly consider the upstream equipment requirements for DGGE (i.e. PCR machines), though I can highly recommend a temperature gradient PCR machine. When applying new primers to an environmental system, having a temperature gradient PCR can be extremely useful, and coupled with DGGE analyses, one can rapidly assess a variety of annealing temperatures.

Before diving into various links to the dominant DGGE manufacturers, I would like to comment that there is a new technology which works in a similar manner, but employs an HPLC column rather than an acrylamide gel for separation. I have not tested this out myself, but the instrument has several theoretical advantages over DGGE analyses, namely that no DGGE gels have to be poured, and that a fraction collector can be employed to recover various “bands”. For those interested, see the following website: www.transgenomic.com/ (denaturing HPLC).

Essentially a DGGE system is a heated fish tank that you can run voltage through. The critical issues for DGGE systems are temperature control and stability of current. There are several major DGGE manufacturers in the market. These include, but are not limited to, the following:

Bio-Rad
http://www.blogger.com/Local%20Settings/Local%20Settings/Temporary%20Internet%20Files/Local%20Settings/Temporary%20Internet%20Files/OLK1B7/www.bio-rad.com
Ingeny
http://www.ingeny.com/
http://www.ingeny.com/ingeny_brochure_eng.pdf
CBS Scientific
http://www.cbssci.com/

I have personal experience with both the Bio-Rad and the Ingeny. I believe that the CBS Scientific is probably the least expensive of the three and the Ingeny the most expensive. The Bio-Rad is probably the most commonly used, and while you can get very nice gels from this machine, there are a number of recurring problems, including:
o poor quality back gels (due to proximity to heating element, lack of adequate stirring)
o easily broken glass plates/spacers/clamps/heating element (all which can be replaced at an expensive cost)
o need to remove the active, heavy part of the DGGE system from the buffer to put gel in, remove gel, add samples, etc (this increases the likelihood of breaking the heating element, and cooling of the buffer during loading and cleaning of gels)
o limited number of lanes and edge smiling (all DGGE systems get this)
o Corrosion of electrodes
o Awful gradient-forming device (I recommend purchasing a different gradient-former).

The Ingeny system, in my experience, has many advantages over the Bio-Rad system, but it also has a few annoying quirks. These advantages and disadvantages:
o No stirring (instead, a re-circulating pump – this also helps with cleaning lanes)
o No need for removal of heating element from buffer tank (this allows heating during cleaning and loading of gel)
o U-shaped spacer eliminates the need for pouring a seal at the bottom of the gel. However, when greasing the spacers (to reduce smiling) the spacers can become warped and are not easy to push down. I have also found that getting rid of bubbles trapped between the bottom of the gel and the “pushed down” U-shaped spacer is the most annoying feature.
o A very large buffer volume (17 L) which probably helps with temperature stability, but is somewhat annoying in terms of buffer preparation.
o Large gels allow 32 lanes (routine) and 48 lanes (very narrow)
o Comes with a reasonable gradient former

I cannot currently comment on the CBS system, but if someone would write in, I would be delighted to add some comments. I have heard that it also has a large buffer volume (18 L) and can hold 4 gels.

In addition to the DGGE machine itself, you will need a:

GRADIENT FORMER

The simplest and most effective design for a gradient former can be purchased from a number of different manufacturers, but I have managed to track down the website location for those from CBS Scientific. The largest gels, as far as I know, are for the Ingeny and have a total volume of about 50-60 ml. These therefore require the CBS Scientific Cat. # GM-100 – which has a total volume of 50 ml on each side.

You will also require a small stir bar (Teflon coated) to stir one chamber of the gradient former, a stir plate, a peristaltic pump (see below), and teflon or similar tubing.

PERISTALTIC PUMPS

http://www.rainin-global.com/rp1.htmlhttp://www.lplc.com/instruments/pump.htmhttp://www.millipore.com/catalogue.nsf/docs/C419

POWER SUPPLY

Any reasonable power supply should serve. It should be able to run 200 V, and up to 300 V may be useful for some users.


PREPARATION OF GELS

Each system will have slightly different preparation methods and you should follow the recommended protocols as best you can. I have attached links to a few web-published protocols. In addition to the equipment listed above, you will need the following items. I highly recommend that you purchase the products already made (e.g. 40% acrylamide solution rather than making your own from powder; buying deionized formamide rather than using a resin , etc):

· A front and back glass plate
· 1 mm spacers (U-shaped spacers for Ingeny system)
· Spacer Grease [I use Dow Corning High Vacuum Grease, but others should be suitable] HIGHLY RECOMMENDED to reduce gel smiling at perimeter. Coat the spacer with a thin layer of grease and wipe off excess grease with a kimwipe.
· Appropriate comb
· Gel casting station
· 40% Acrylamide (37.5: 1, acrylamide:bis-acrylamide)
· Formamide (deionized)
· Urea
· 50X TAE (50X is 242g Tris base; 57.1 ml glacial acetic acid; 100 ml 0.5M EDTA (pH 8) per liter)
· Ammonium Persulfate [Is highly hygroscopic – remember to buy fresh routinely]
· TEMED (N,N,N',N' -tetramethylenediamine)
· Nucleic acid stain
o GelStar
o EvaGreen
o Ethidium Bromide [Not recommended due to low sensitivity, toxicity]

o Prior to casting of the gels, you will need to have made stock solutions of a zero denaturant acrylamide solution and a high denaturant acrylamide solution (I generally make 80% solutions, defined below).

The zero denaturant acrylamide solution will vary slightly according to final acrylamide solution, but a 100 ml stock solution of 6% acrylamide, zero denaturant solution will contain 15 ml of 40% acrylamide (37.5: 1, acrylamide:bis-acrylamide), 2 ml of 50X TAE, and water to 100 ml. A 100 ml stock solution of 6% acrylamide, 80% denaturant will contain 15 ml of 40% acrylamide, 33.6g of Urea, 32 ml of formamide, 2 ml of 50X TAE and water to 100 ml. This can be a bit tricky to make. I usually add all the liquids together first (acrylamide, formamide, TAE and a bit of water) and then add the Urea. This can take a while to get into solution and mild heating can help, though it shouldn’t be necessary. Not to be repetitive, but many of these compounds are highly toxic. Higher acrylamide concentrations can be even trickier when making the high denaturant solution – be careful not to add too much water.

Prior to casting of the gel, specific concentrations of denaturing solutions can be made by mixing the zero and high denaturant solutions in appropriate ratios. I prefer to make the solutions by mixing integer values of each solution (therefore, no 6.35 ml etc). This will get you reasonably close to the value you want (for example, for the Ingeny system I used 16 ml of zero denaturant and 6 ml of 80% denaturant solutions to get a 22% solution). Since you should be pouring a gradient that is wider than the region you want, a little variation at either extreme is not particularly significant.

I make fresh APS (10% solution, in water) fresh before each gel. However, aliquots can be frozen and used for some time (ca. 1-2 wks). In general, fresher reagents are better and a number of problems with DGGE gels can be traced to old acrylamide, formamide and TEMED solutions. These solutions are certainly a place to start when trying to trouble shoot. Nonetheless, I have found that acrylamide solutions, kept at 4C, can keep for months without any negative effect. Well cleaned glass plates also help improve gel quality. Some DGGE operators clean the glass plates with ethanol as a last stage; I find that this often diminishes the quality of the gel pouring.

Web Protocols (For Bio-Rad)

Protocol from the Laboratory for Microbial Ecology Department of Earth, Ecological and Environmental Sciences University of Toledo
http://www.eeescience.utoledo.edu/Faculty/Sigler/RESEARCH/Protocols/DGGE/DGGE.pdf

G. Zwart and J. Bok, dept. of Microbial Ecology, Center for Limnology, Netherlands Institute of Ecology (NIOO-KNAW). E-mail: http://www.blogger.com/Local%20Settings/Local%20Settings/Temporary%20Internet%20Files/Local%20Settings/Temporary%20Internet%20Files/OLK1B7/Zwart@cl.nioo.knaw.nl
http://www.kuleuven.ac.be/bio/eco/bioman/publications/Protocol-DGGE_bacteria&protists.pdf

Well, I haven’t found any online protocols for Ingeny or CBS (I haven’t looked very hard), but pouring these gels is essentially the same as any other gel.

PCR / NESTED PCR FOR DGGE

DGGE standards
There are no established standards for DGGE. However, applying homemade standards to DGGE gels can be very useful. For some gel analyses programs, it is useful, if not essential, to have standards every few lanes or so due to the heterogeneity of the DGGE gels. I have seen some users load size standards onto DGGE gels, though these don’t have much meaning. The best approach is to create your own set of standards by mixing PCR product of a number of differently migrating clones. You should run the PCR of each clone independently and then mix the PCR yield. Make a large stock.

In addition, you may want to check out a new technique of adding standards to each lane. This can be found in Section 8.0 Gel Analysis. See the article by Neufeld and Mohn, 2005.

PCR product cleanup prior to DGGE analyses
There has been some discussion in the Yahoo DGGE group as to whether or not it is useful to cleanup PCR products prior to DGGE analysis. I have never routinely performed PCR cleanups and I think it would be a major added expense and effort. Luckily, there doesn’t seem to be any major reason to cleanup the PCR product. The primers and primer dimers migrate out of the gel and do not interfere with final staining analyses. However, you may need to cleanup the PCR product if you want to do cloning or quantification.

How much PCR product to load on DGGE?
There is no established amount of PCR product to load on DGGE. However, some things to consider: the lower the diversity your system, the less DNA you need to load. For example, if you load 50 uL of DNA from a PCR of a single clone, you will definitely load too much DNA. However, in a highly diverse microbial community, that PCR product will be split among a large number of different sequences, making detection of some problematic. Another rule of thumb suggests that if (ignoring for the moment any issues with PCR bias) a sequence is not at least 1% of the total population targeted by the primer set, that sequence will be difficult to detect by DGGE. Thus, specific primers can aid in detecting less abundant populations by narrowing the range of target population.

From the DGGE group I have seen suggestions that 300-500 ng of DNA is an appropriate amount to load per lane – for an environmental sample, or more specifically, 2-20 ng per band, when using non-EtBr stains. Obviously, for PCR product of a pure culture or clone, you’ll need significantly less. Although not widely done, it probably would be a good idea to quantify PCR products prior to loading on DGGE gels, so as to be able to compare intensities between lanes. If you have weak PCR, you can load large volume into each lane by loading twice. That is, fill the well, electrophorese for a while and the load the rest of the PCR product. Since DNA fragments often stop when they reach their specific denaturant concentration, and run times are probably longer than necessary, there does not appear to be any negative effect resulting from sequential DNA addition.

One thought – the size of the PCR fragment may also be a consideration. Given the same copy number, a longer DNA fragment will yield a stronger fluorescent signal.

How much DNA to put in each PCR reaction?
This topic has come up recently in the Yahoo group. Ideally, a similar amount of DNA should be added to each PCR reaction, but in reality, this is often time consuming to do. This requires the measurement of DNA concentration from each DNA extract, and then dilution of each DNA to the same final concentration. This may be particularly important for quantitative PCR. For non-quantitative PCR of environmental samples ultimately used for DGGE, there seems to be a range of 10 to 100 ng (per 50 microliter reaction) in use. I have seen on the DGGE Yahoo group that some users have been diluting genomic DNA prior to adding it to PCR. Diluting DNA to avoid contaminants such as humic acids/polysaccharides may work sometimes, but in general, it is probably better to clean up the DNA, even if you loose a proportion of your DNA. The article below suggests that diluting DNA makes reproducibility worse.
· Chandler DP, Fredrickson JK, and Brockman FJ. 1997. Effect of PCR template concentration on the composition and distribution of total community 16S rDNA clone libraries. Molecular Ecology 6: 475-482

Some situations can be far more complex. As Kim discusses in post #1267, one can have a situation where there is a variable amount of non-target genomic DNA. Thus, it may not be entirely fair to add the same total amount (ng) of genomic DNA to each PCR reaction, if the amount of target DNA varies from sample to sample. The example given is that of DNA extracts from plant samples. If one is looking at microbial communities associated with plant roots, one might recover variable amounts of root DNA depending on growth stage, etc. I don’t have a specific answer to the question of whether or not one should adjust the amount of genomic DNA added to PCR reactions based on the knowledge that variable amounts of non-target DNA are present. It seems likely that in many environments there will be variable amounts of non-target DNA (e.g. fungal DNA in bacterial population analyses, etc.) and that these cannot be readily quantified. Since standard PCR is not overly sensitive to starting concentration (thus the requirement for real-time PCR) this may not be something to worry about too much.

Primer dimerization
Some PCR primers form strong primer dimers than can be visualized by agarose electrophoresis [See figure below]. I have not found that these interfere with DGGE analyses as they appear to migrate out of the gel. However, these primer dimmers can be a problem if you run ligation/transformation/sequencing reactions directly from the PCR product. The high quantity of the primer dimer can decrease the efficiency of the cloning reaction because the dimer will compete for plasmid. This is because the primer dimers are double-stranded, and are A-tailed (if you are using Taq polymerase) just as the PCR product of the correct size. Since these primer dimers can be extremely abundant, and are double-stranded and a-tailed, they can compete with the primary PCR product in the ligation reaction. Thus, many of the clones will contain only the primer dimers, decreasing the efficiency of the ligation/transformation reaction. Thus, you may want to clean up your PCR product prior to cloning. Remember, if a band of smaller size (bp) is of the same intensity as a band of larger size, there are actually a greater number of copies of the smaller fragment.

Nested PCR
Nested PCR is a technique by which genomic DNA is subject to PCR amplification and then the product resulting from the amplification is subject to a second, nested PCR with primers that target a region within the region targeted by the first PCR primers. There are a number of reasons to employ nested PCR:
· Increase PCR yield from weak reactions
· Avoid detrimental effects of PCR amplification with primers that have a GC-clamp
· Generate specific DNA fragments suitable for DGGE analysis from DNA fragments that are not suitable for DGGE analysis
· Allow direct DGGE comparisons of general and specific population analyses
· Recover additional sequence information than that provided in DGGE-appropriate fragments.
· Avoid having to develop and optimize new DGGE conditions for a new primer set.
· Compare the community recovered by different primer sets (for example - PCR large fragments with different general bacterial primer sets and then nest with the same general bacterial primer sets and compare by DGGE. This is limited by the internal primer set, but it can still reveal important differences in primer sets.)

I have added a small flow chart below to show some of the potential. I think that in particular, the capacity to generate sequence information that is larger than the DGGE fragment is underutilized.

There are some caveats related to nested PCR. Namely, since there are multiple PCR reactions and high numbers of PCR cycles, the potential for chimera formation and sequence errors increase. Also, you will find that nested PCR can often generate secondary, non-specific bands. You should optimize PCR conditions (temp, cycle #, magnesium concentration) to reduce the intensity of these bands, if possible. All things being equal, if you can achieve your aims without nested PCR, you probably should avoid it. However, for some things, the nested PCR approach is so useful, it’s worth it. If you are worried about the quality of sequences recovered from nested PCR, you can apply specific primers (developed based on the sequence information recovered from the nested PCR approach) to the same samples to demonstrate that the sequences are actually present.

When I conduct nested PCR, I usually dilute the original PCR product 1:5 or 1:10. I do not perform a cleanup prior to the second stage of PCR. However, I generally raise the annealing temperature of the second stage of PCR by 4C relative to the standard temperature for the given primer set (this will also help avoid contamination of the blank, by the way…), and I perform fewer cycles and often I add less primers. These are all attempts to limit the efficiency of the PCR reaction so as to not get too many secondary, non-specific bands.

· Shabir A. Dar, J. Gijs Kuenen, and Gerard Muyzer. 2005. Nested PCR-Denaturing Gradient Gel Electrophoresis Approach To Determine the Diversity of Sulfate-Reducing Bacteria in Complex Microbial Communities. Appl. Environ. Microbiol. 71:2325-2330.
· Wood GS, Uluer AZ. 1999. Polymerase chain reaction/denaturing gradient gel electrophoresis (PCR/DGGE): sensitivity, band pattern analysis, and methodologic optimization. Am J Dermatopathol. 1999 Dec;21(6):547-51.
· Green, S.J., Freeman, S., Hadar, Y. and Minz, D. 2004. Molecular tools for isolate and community studies of Pyrenomycete fungi. Mycologia, 96(3), 2004, pp. 439-451.


ELECTROPHORESIS

· Vanessa M. Hayes, Ying Wu, Jan Osinga, Inge M. Mulder, Pieter van der Vlies, Peter Elfferich, Charles H. C. M. Buys, Robert M. W. Hofstra. 1999. Improvements in gel composition and electrophoretic conditions for broad-range mutation analysis by denaturing gradient gel electrophoresis. Nucleic Acids Research. 27(20): 29.

RUN TIME / VOLTAGE
There has been considerable discussion on the yahoo group about the correct run times and voltages. In general, longer run times with lower voltages tend to produce better quality gels. Voltages are in the range of 50-250V, and run times are generally from 3 hr to 17 hr. A standard run time of 17 hr (essentially overnight) at 100 V produces high quality gels for many PCR fragments. The exact voltage may differ between PCR product size, buffer concentration, buffer temperature, run time, and gel acrylamide percentage. For very short fragments, it may be important to use relatively low voltages for a long run to avoid having the fragments migrate out of the bottom of the gel. For gels in which the fragments completely stop within the gradient, extra run time will probably not effect the final gel. It seems that Volt-Hours are probably the best thing to consider. We used to run some gels at 250 V for 3 hr and found 17 hrs at 100 V yielded better gels; the 100V run had 1700 volt-hrs while the 250 V run had only 750. It should be noted that if you are particularly worried about running your fragments out of the gel, there may be a problem with your gradient (i.e. not wide enough) or your acrylamide concentration (not high enough). Every user will have a preferred voltage and run time, so I am not going to make any absolute recommendations here – there appear to be many possible conditions which yield good results. Many times the conditions will have to be empirically determined.

INITIAL VOLTAGE VALUES
Some users will run a high voltage initially to bring the DNA into the gel and then use a lower voltage for the rest of the run. In the Ingeny system, one is supposed to run the gel for a little while to draw the DNA into the gel before turning on the recirculating pump – otherwise the pump may stir up the PCR product in each well. I do not think that running an initial high voltage is important; the critical factor is to have the temperature stabilized. With the Bio-Rad, you have to keep removing the heating element from the tank and during the cleaning of the wells and loading of samples, the buffer cools down. Before starting the electrophoresis, you should make sure that the temperature of the tank is at 60C (or whatever temperature you are using). You can run a low voltage while the tank is heating to its final temperature just so you limit diffusion of the DNA into adjacent wells. The important thing is to not let the DNA migrate into the gradient until the buffer is at the correct temperature.

AMPERES VS VOLTS
I have seen a lot of discussion about voltage vs amperage when dealing with electrophoresis. You could run your electrophoresis with either fixed voltage or amperage, but the standard appears to be constant voltage. I have found that when running gels with a fixed voltage, the resulting amperage is an excellent indicator for problems with the system. Old buffer, leaks in the system, air bubble blocking the bottom of the gel, poor circulation, temperature – all these factors can affect the operation of the system and these will show up in the amperage – usually by low amperage relative to a well operating system. For example, on my Ingeny DGGE system, I can expect approximately 40 mA when running a single gel at 100V, 60C. To make sure my system is properly set up prior to loading samples, I run a few minutes of electricity through the system at 100V to make sure I get 40 mA (obviously each system will have a different “standard” mA value at a fixed voltage). If the values are way off I know there is some sort of problem. I never load my samples until the mA values are within proper ranges. You should also remember that at the same voltage, running 2 gels will yield a higher mA than running 1 gel. Regardless of how many gels you are running, you should maintain the same voltage. If your gel box cannot handle the amperage from multiple gels, you may have to run at a lower voltage for a longer time.

GEL STAINING AND VISUALIZATION
Do NOT add stain to the gel prior to electrophoresis. Staining of DGGE gels MUST be done after electrophoresis. After stopping the electrophoresis and having separated the gel plates, leaving the gel attached to one plate, you are ready to stain the gel. There are a number of nucleic acid dyes which are adequate for visualization of DGGE gels. I recommend not using Ethidium Bromide (EtBr). EtBr is a strong mutagen and has a much lower sensitivity that some of newer nucleic acid dyes available (listed above). EtBr also cannot be excited well by wavelengths above 400 nm. This is a major downside if you want to use non-UV illumination tables (see below).

I am currently using GelStar as a nucleic acid stain. It is somewhat expensive, but not prohibitively so, is less toxic and more sensitive that EtBr. I recommend staining while shaking for 30 minutes and then transferring to a de-staining tank (water is fine) for another 15-30 minutes. GelStar and similar stains will be useful if you want to use a non-UV illumination table. I can recommend the Dark Reader illumination table : http://www.clarechemical.com/. This product excites the nucleic acid dyes at wavelengths from roughly 400 – 500 nm. Then a second filter is placed over the gel which blocks out light below 500 nm. Since the dyes emit at wavelengths above 500 nm, the gels can be visualized. The advantages of this table are that there is no UV light, and no damage to the DNA in the gels. If you are intending to excise bands from DGGE gels, I can recommend this table. There is no hurry to excise bands without doing damage to the DNA in your gel or in your body, and you can place the gel, with the glass plate, onto the table. The quality of the pictures from this table is not as good as a UV table. Thus, for publication quality photos, you’d better have a UV transilluminator. The Dark Reader also comes with goggles that contain the second filter. Thus you can easily excise bands without any filter in the way.

SILVER STAINING
I’m not sure anybody still silver stains, but here’s a reference. I remember some users had problems extracting bands and re-PCRing after silver staining. One advantage is that the signal does not degrade rapidly as with some of the newer stains.
· D. Radojkovica and J. Kusic. 2000. Silver Staining of Denaturing Gradient Gel Electrophoresis Gels. Clinical Chemistry 46: 883-884.

GEL TRANSFER FROM BUFFER TO UV TABLE
Transferring the gels from the staining bath to the UV table can be a tricky and annoying procedure. I have seen published papers with 5 or 6 DGGE gels and every single one of the gels is ripped somewhere. Handling the thin gels, particularly the 6% acrylamide gels, takes experience. I previously posted some tips (see below) on the Yahoo DGGE group, and the best tip is to keep everything lubricated with buffer or water – this reduces friction and tearing.

The best word is patience. Do everything slowly and make sure there is plenty of liquid around to lubricate the transfer of the gel. You can reduce your anxiety by removing one step - when you transfer the gel to staining, keep the gel on the bottom plate and simply put the whole plate in the staining bath. I shake the gel off the bottom plate while it is in the staining bath to make sure that the dye can diffuse in from all directions. When lifting the gel out of the staining bath (and then into a de-staining bath and/or directly onto the UV table) lift the plate out of the staining buffer partially - this allows you to see where the gel is. If the gel is centered on the plate, place one palm on top of the gel to keep it in place while you lift the whole plate out (with the gel) using the other hand. Let some of the liquid drain off the plate (keep your palm on the gel). Now place the plate on your UV table at an angle and let the gel slide onto the table (gently, slowly, patience....); as the gel comes off slide the plate further away. Have a water bottle available and add water when needed. Also, you may have to nudge the gel if it gets stuck - particularly at the edges where there is grease (from the spacers). Don't worry about having the gel be perfect as it slides off the glass plate. It can be adjusted easily on the UV table with gentle nudging and more water.

ADD GLYCEROL TO INCREASE GEL FLEXIBILITY
Dr. Von Sigler of the Laboratory for Microbial Ecology (University of Toledo) has this to say about glycerol: “ I add glycerol to a final concentration of 2% (v/v) because it adds a great amount of flexibility to the gels. This added flexibility comes in handy when manipulating the gels (removal from the plates, staining, placement and positioning on transilluminator) and decreases the risk of tearing the gel. It does not impact DNA migration or the banding qualities.”

GEL ANALYSIS
I cannot claim to be a great expert on dendogram-style analyses of DGGE gels. I am going to list some known programs, some articles and hope for some contributions from more knowledgeable users. Some pitfalls to be considered, however, are the problem of multiple populations co-migrating to a single band position, multiple bands from a single organism, and PCR artifacts resulting from degenerate PCR primers. In addition, if the dendogram analyses also consider the intensity of each band, then it should be important to either load the same amount of DNA in each lane, or to make intensity values relative to total intensity. Some of the commercially available programs can be extremely expensive.

I highly recommend a new manuscript in AEM by Josh Neufeld and William W. Mohn (“Fluorophore-labeled primers improve the sensitivity, versatility, and normalization of denaturing gradient gel electrophoresis”). This manuscript develops a new technique to increase our capacity to use DGGE gels for digital analysis by including standards within each lane. These standards have fluorescent tags that fluoresce at different wavelengths than the fluorescent molecules attached to the unknown PCR product. Brilliant! The caveat is as the authors note, besides expense, “that access to an expensive laser-scanning instrument is required, which may limit widespread use of this application at this time.” One other thing: since the sensitivity is increased by this method, the authors could reduce the number of PCR cycles required to generate enough DNA for DGGE. This is another advantage.

· Neufeld, JD and WW Mohn. 2005. Fluorophore-labeled primers improve the sensitivity, versatility, and normalization of denaturing gradient gel electrophoresis. Appl. Environ. Microbiol. 71:4893-4896.

Jaak Truu of the Yahoo DGGE group has this to say: “Standard ordination methods (PCA; CA) do not take into account the situation that one band may represent several species. There are some similarities between data obtained with DGGE and microarrays (noisy data). In case of microarray data new methods and software applications are developing very fast. For DGGE there are some papers considering in depth statistical analysis of microbial community fingerprints (Wilbur et al., 2002 [See Below]). The simplest way (but not only one) to confirm the grouping or clustering of your DGGE data obtained with cluster analysis or ordination is use methods that implement bootstrapping, randomization or MonteCarlo methods. Unfortunately these methods are not included in generally used statistical analysis software.”

· Tong Zhang and Herbert H.P. Fang. 2000. Digitization of DGGE (denaturing gradient gel electrophoresis) profile and cluster analysis of microbial communities. Biotechnology Letters. 22: 399 – 405.
· Fromin, N., Hamelin, J., Tarnawski, S., Roesti, D., Jourdain-Miserez, K., Forestier, N., Teyssier-Cuvelle, S., Gillet, F., Aragno, M. & Rossi, P. 2002. Statistical analysis of denaturing gel electrophoresis (DGE) fingerprinting patterns. Environmental Microbiology 4 (11), 634-643.
· J. Wilbur, J.K. Ghosh, C.H. Nakatsu, S.M. Brouder, and R.W. Doerge. 2002. Variable selection in high-dimensional multivariate binary data with application to the analysis of microbial community DNA fingerprints. Biometrics 58:378-386.

GEL ANALYSIS PRODUCTS
· BioNumerics (Applied Maths, Kortrijk, Belgium)
· Quantity One (Bio-Rad)
· DGGESTAT
· Gelcompare II
· Bioprofil (Vilbert-Lourmat)
· Jayson D. Wilbur from Purdue University has written “There is a free statistical software package called "R" that can be used to construct dendograms based on the Ward method. It is available at: http://www.r-project.org/ You will also need to download the "cluster" package from the website and use the commands "agnes" and "plot.agnes" (specifying the option method="ward").”

BAND EXCISION AND ANALYSIS
This subject is probably one of the most frequently discussed on the Yahoo DGGE group. Having done many, many band excisions, I cannot say that I find it particularly enjoyable and I believe, as I mentioned on the Yahoo Group (see below), that it is not the best way to go about acquiring sequences. However, for bands of particular interest that cannot be acquired in any other manner, it should be used.

Avoiding Band Excision by Direct PCR product Sequencing
The choice of bands to excise may also be highly subjective, and, as many of us have experienced, there can be multiple sequences hiding in a single band. Thus, excising, cloning, and sequencing of a single clone from each band can miss hidden diversity. Furthermore, excision of bands is time consuming, and if done on a UV table, unpleasant and can be damaging to DNA (yours and your sample's). If you want to be sure that the excised band is the correct thing, you should re-PCR, run another DGGE, clone the PCR product, and then screen the clones with another DGGE analysis. All of this work makes me wonder - what is the point? I would like to suggest that a better way to approach this is to clone directly from the original PCR product (using GC-clamped PCR primers) and then screen the clones against the environmental sample. Thus, from a single cloning reaction you will be able to pick off many of the dominant bands and you will only have to run a single DGGE instead of 3. I would suggest that excision of bands should only be done in those cases where important bands simply cannot be recovered by an initial cloning reaction. In addition, at least 2 clones for each band position should be sequenced - this can help verify if there is hidden diversity at each band position.All of this cloning would have been an onerous burden. However, there are now sequencing facilities that will take clones in 96-well plates, extract plasmid and run sequencing reactions for approximately $3/rxn [NOTE: this is in the year 2005]. This means that you can avoid having to do plasmid extractions yourself and that the cost will still be less than you used to pay. So, perhaps we should pick more colonies, clone more and excise bands less.

High-throughput sequencing techniques

· Neufeld, JD, Yu, Z, Lam, W, and WW Mohn. 2004. Serial Analysis of Ribosomal Sequence Tags (SARST): a high-throughput method for profiling complex microbial communities. Environ. Microbiol. 6:131-144.
· Kysela, David T., Palacios, Carmen & Sogin, Mitchell L. 2005. Serial analysis of V6 ribosomal sequence tags (SARST-V6): a method for efficient, high-throughput analysis of microbial community composition. Environmental Microbiology 7 (3), 356-364.
· Yu, Zhongtang, Yu, Marie & Morrison, Mark. 2005. Improved serial analysis of V1 ribosomal sequence tags (SARST-V1) provides a rapid, comprehensive, sequence-based characterization of bacterial diversity and community composition. Environmental Microbiology

HOW TO EXCISE DGGE BANDS
You can excise DGGE bands with two basic approaches. The first is to take a sterile pipette tip by hand and stab it into the gel at the position of the band that you are interested in. Upon removing the tip, you then inoculate a fresh PCR reaction with the DNA adhering to the tip. I have had very mixed results with this method and do not recommend it. While it is certainly time saving, in a sense, you often do not get the band that you are interested in.

The second technique is to physically excise the band with a sterile razor blade. You should be careful to limit your exposure to UV (if you are using a UV transilluminator), and there is a limit to the amount of exposure your DGGE gel can get before the DNA is significantly damaged and not particularly usable for downstream analyses. Both these techniques can be difficult when the bands are very close together. In all cases, it is critical that the PCR product from this re-amplification be screened against the environmental sample on a second DGGE analyses to ensure that the PCR product reflects the excised band.

WHAT TO DO WITH YOUR EXCISED BAND
Bead-beating
Place the acrylamide fragment in a tube with some glass beads and water (or TE) and bead-beat or vortex briefly. Then incubate the tube either at 37C for 30 min, or 4C for several hours/overnight. A microliter or two of the water can then be used as a template for a subsequent PCR reaction. I would centrifuge the tube prior to taking liquid for the PCR reaction to remove any possible acrylamide pieces. Remember not to overload the PCR reaction with DNA – this can cause subsequent problems with smearing on DGGE gels.

Gel electrophoresis and cleanup
This is a clever technique to transfer the DNA in the acrylamide to agarose. The agarose can then be melted and the DNA recovered. Place the acrylamide fragment into a well in an agarose gel and fill the well with fresh liquid agarose. Allow the agarose to cool and gel, and then electrophorese the gel. The DNA will migrate out of the acrylamide and into the agarose gel. The fragment can then be excised from agarose and cleaned up either with a DNA gel cleanup kit or a sodium iodide/silica cleanup (see below):
· Boyle JS, Lew AM. 1995. An inexpensive alternative to glassmilk for DNA purification. Trends in Genetics. 11(1):8.

If you re-PCR the excised DNA, you will definitely need to screen the PCR product against the original environmental sample with a second DGGE. Alternatively, you can clone directly, and then screen the clones against the environmental sample. Under no conditions is it a good idea to clone directly and send for sequencing without verifying that the clone band is reflective of the environmental band that you were interested in.

You may want to use one of your PCR primers for sequencing analyses, even if you have cloned your PCR product. I recommend using the primer without the GC-clamp. In that manner, the sequencing reaction begins at the end of the fragment without the GC clamp, and you don’t waste the best stage of the sequencing reaction on the GC clamp region. If your fragment is short enough, you probably don’t have to worry.

WHAT TO DO IF YOUR EXCISED BAND ISN’T PURE
This is a common enough problem that has no particularly easy solution. When excising bands, or stabbing bands into PCR, it is easy enough to pick up unwanted DNA. Thus, screening of excised bands, cloning, and screening of cloned bands are all part of the work required to ensure that you have isolated the correct band. Even if an excised band isn’t pure –that it, a mixture of several bands, it can often be much less diverse than the original sample and can help you isolate the band of interest. Remember, if you are going to clone the excised band (highly recommended), you simply need to get the ratio of target:non-target high enough to make it likely that you will recover the band of interest by cloning.

A PROBLEMS CHECKLIST

Ø Back gel on Bio-Rad DGGE is often of poor quality perhaps due to inadequate mixing and proximity to the heating element.
Ø Are you using the Bio-Rad wheel to cast gels? Get yourself a gradient former (see above).
Ø Did you clean the wells properly? Unpolymerized acrylamide and urea and formamide diffusing upwards can cause problems. Clean wells thoroughly before loading.
Ø Did you grease the spacers? This can help reduce smiling on the edges.
Ø Are there air bubbles in your gel? Try to avoid air bubbles getting trapped in your gel during pouring. Tap on the glass to cause the bubbles to rise to the surface. Bubbles in gels can cause streaking in the lanes. Avoid air bubbles getting trapped under the comb by inserting the comb at an angle and slowly.
Ø Are your reagents fresh? Many problems have been traced to old or poor quality reagents both for DGGE and for PCR. dNTPs, acrylamide, and formamide have repeatedly come up as the culprit in poor quality DGGEs on the Yahoo DGGE Group.
Ø Are your reagents high quality? Molecular Biology grade chemicals should be used.

TROUBLESHOOTING AND ADVANCED TOPICS

ACRYLAMIDE PERCENTAGE
You will have to determine what acrylamide solution is correct for your system and it may vary according to primer set. For most environmental DGGE analyses 6% acrylamide is used. Shorter fragments may use an 8% acrylamide solution and in some cases an acrylamide gradient can also be included. I have used acrylamide solutions of up to 12% (rare) [See below].
· Green, S.J., Freeman, S., Hadar, Y. and Minz, D. 2004. Molecular tools for isolate and community studies of Pyrenomycete fungi. Mycologia, 96(3), 2004, pp. 439-451.

I have found with one primer set generating a PCR product of about 400 bp, that an 8% gel gave much worse results that a 6% gel. I was really surprised – I had been running the primer set at 8% acrylamide for a long time and had reasonable results, but limited visible diversity. By chance I ran the sample PCR product on a 6% gel and got amazingly higher diversity and better separation. I think ultimately empirical tests are required when optimizing any new primer set for DGGE analysis. However, it seems reasonable to think that small fragments would be better off with higher acrylamide percentages.

BLURRY, FUZZY OR SMEARED BANDS / SUDDEN CHANGE IN QUALITY OF GELS
These types of problems are often a sign of PCR problems. However, they can also indicate a poorly made gel, poor quality or old reagents, an improper gradient, irregular current, old buffer, temperature control issues (e.g. back-gel issues on the Bio-Rad), etc. Also, you may want to consider if your reagents, even if fresh, are really good. I have been satisfied with Bio-Rad reagents and Sigma reagents as well. It is best to be consistent once you have found good reagents.

If your problem can be traced to using the back gel on the Bio-Rad machine, you have a couple possible solutions:
>> Don’t use the back gel
>> Place the entire apparatus in a water bath heated to the same operating temperature. This will help the system maintain temperature control better.
>> Place a stir bar in the bottom of the tank and put the entire apparatus on a stir plate to increase mixing of the system. Be careful – some stir plates cannot handle the weight from the full Bio-Rad system (This is from personal experience….).

Bands in the top of the gel are often fuzzy and indistinct, and this may be an indication that these are some sort of artifact of the PCR/DGGE analysis. In particular, if I notice a band that is present in all samples, very high in the gel, and fuzzy, I am highly suspicious of it. These can be heteroduplex bands which denature rapidly, or perhaps single stranded DNA.

For sudden change in gel quality:
>> Check primer quality; re-order primers if old
>> Check age of all DGGE reagents: acrylamide, formamide, APS, TEMED.
>> Are you using new reagents in PCR? Check if dNTPs, Taq, etc. are good.

BUFFER RE-USE
There is some discussion of how many runs can be done on a single tank of buffer before replacing the buffer is required. In my experience, there is a maximum of 4-5 runs per tank. The Ingeny system recommends that you replace 5L of buffer every run and replace the entire tank (17L) every 3 runs. I have had no problem running 4 gels without replacing the buffer at all on the Ingeny. It is advisable, however, to use fresh buffer if the gel is to be for a publication.

In addition, if you run your buffer at 0.5X TAE, it may be necessary to change buffer more frequently than with a 1.0X TAE buffer strength.

Detection issues / DGGE SENSITIVITY
· Li Zhang, Stephen Danon, Martin Grehan, Adrian Lee, and Hazel Mitchell. 2005. Template DNA Ratio can Affect Detection by Genus-Specific PCR–Denaturing Gradient Gel Electrophoresis of Bacteria Present at Low Abundance in Mixed Populations. Helicobacter. Volume 10 Issue 1 Page 80.
· Trulzsch B, Krohn K, Wonerow P, Paschke R. 1999. DGGE is more sensitive for the detection of somatic point mutations than direct sequencing. Biotechniques. 27(2):266-8.

DGGE ANALYSIS OF CLONES
In some cases, PCR conditions optimized for environmental samples do not work well for PCR amplification of clone DNA. In part, I suspect this is due to the exceedingly high concentration of DNA added to the PCR reaction from a clone (can be as cellular material or boiled cellular material or from plasmid extraction) and the high copy number of identical sequence. I recommend a higher dilution of DNA, fewer cycles, lower magnesium concentration, and higher annealing temperature – something to reduce this problem when amplifying clones. I have noticed that in general, pure culture DNA can behave differently in PCR reactions that in mixed cultures or environmental samples. This can make optimization of new primer sets difficult. One could spike environmental samples with target DNA and perform optimization thusly to avoid such a problem.

· S.A. Middleton, G. Anzenberger, and L.A. Knapp. 2004. Denaturing gradient gel electrophoresis (DGGE) screening of clones prior to sequencing. Molecular Ecology Notes 4 (4), 776-778.

Darek Bulinski has this to say about analyzing multiple clones per lane: “If we have a lot
to screen we run ten, 5 microliter samples per lane. This allows us to look at a large number of samples per DGGE gel. If any given lane contains band(s) that correspond to our whole community sample, we then run the samples in individual lanes on a second gel to identify the clone from which it was amplified. Also, for running clones on a DGGE gel, we pick a colony into 100 microliters of PCR water, vortex it and use 1 microliter per 25 microliter reaction (we've never had problems with this or required boiling). Then running 5 microliters of that on a DGGE gel is plenty (more if the amplification was less efficient).”

GC CLAMPS
Remember to put your GC clamp at the 5’ end of the primer!

· RM Myers, SG Fischer, T Maniatis and LS Lerman. 1985. Modification of the melting properties of duplex DNA by attachment of a GC-rich DNA sequence as determined by denaturing gradient gel electrophoresis. Nucleic Acids Res. 13, 3111-3129.
· RM Myers, SG Fischer, LS Lerman, and T Maniatis. 1985. Nearly all single base substitutions in DNA fragments joined to a GC- clamp can be detected by denaturing gradient gel electrophoresis. Nucleic Acids Res. 13: 3131 - 3145.

I just happened across a website suggesting that having GC clamps on both PCR primers yielded better separation. I have had poor luck when I accidentally ordered GC clamps for both primers, but it appears this can vary according to the primer set. [See “Bipolar clamping versus monopolar clamping” on this website: http://www.charite.de/bioinf/tgge/ ]

HIGH DIVERSITY ENVIRONMENTS – MEASURING DIVERSITY
Many users have had problems performing DGGE analyses on high diversity environments such as soils. When applying general bacterial primers to the systems, with subsequent DGGE analysis, the gels can look smeared, in part due to the very high number of bands (many of the weak and indistinct). Molecular analyses such as DGGE, TGGE, TRFLP, SSCP can all be troublesome with such high diversity. In such cases, the best approach is probably to build a clone library from the PCR product and generate diversity estimates using rarefaction type analyses (see DOTUR, below). DGGE really reaches its limitations when dealing with such high diversity limitations. I would recommend to anyone dealing with such a high diversity to design narrower primers to apply to the system. Thus, one can focus on, for example, Actinomycetes, for which there are highly specific primers. You can use those primers directly for DGGE or nest them with general bacterial primers.

· DOTUR: http://www.plantpath.wisc.edu/fac/joh/DOTUR/documentation.html
· Schloss, P.D. & Handelsman, J. 2005. Introducing DOTUR, a computer program for defining operational taxonomic units and estimating species richness. Applied and Environmental Microbiology. 71(3):1501-1506.
· Huges JB, Hellmann JJ, Ricketts TH, Bohannan BJM. 2001. Counting the Uncoutable: Statistical Approaches to Estimating Microbial Diversity. Applied and Environmental Microbiology 67 (10) 4399-4406.
· Curtis TP, Sloan WT. 2004. Prokaryotic diversity and its limits: microbial community structure in nature and implications for microbial ecology. CURRENT OPINION IN MICROBIOLOGY 7 (3): 221-226
· William T. Sloan and Jack W. Scannell. 2002. Estimating prokaryotic diversity and its limits. PNAS 99 (16): 10494-10499.

MIGRATION OF BANDS INTO THE GEL IS LIMITED OR ABSENT? (NO VISIBLE BANDS IS A FREQUENT COMPLAINT)
Things to check:
>> Are voltage/amperage levels appropriate?
>> Are there bubbles underneath the gel (Ingeny)
>> Did you pour the gel correctly (i.e. put the high and low acrylamide solutions in the correct wells)?
>> Did you turn on the power supply (this just happened to somebody I know…!).
>> Are the electrode terminals clean (Bio-Rad)?
>> If you have only one gel, do you have a back glass plate to enclose the upper buffer reservoir?
>> Is there a leak in the upper buffer reservoir?
>> Make sure the lid is securely closed (Bio-Rad).
>> Is your stain fresh? GelStar, SybrGreen, etc. all are light sensitive and degrade.
>> Do you have a strong PCR product? Have you added enough DNA?
>> Did you electrophorese for too long (or too high a voltage) and the fragments migrated out of the gel?
>> Did you hook up the electrodes in the right order?
>> Did you let the product diffuse out of the wells in the gel?
>> Did recirculating buffer mix the PCR product in each well?

MULTIPLE BANDING FROM SINGLE POPULATIONS AND MULTIPLE SEQUENCES FROM SINGLE POPULATIONS

· Klappenbach JA, Dunbar JM, Schmidt TM. 2000. rRNA operon copy number reflects ecological strategies of bacteria. Appl. Environ. Microbiol. 66:1328-33.
· S.P. Gafan and D.A. Spratt. 2005. Denaturing gradient gel electrophoresis gel expansion (DGGEGE) - An attempt to resolve the limitations of co-migration in the DGGE of complex polymicrobial communities. FEMS Microbiol Lett. 2005
· A. Schmalenberger and C.C. Tebbe. 2003. Bacterial diversity in maize rhizospheres: conclusions on the use of genetic profiles based on PCR-amplified partial small subunit rRNA genes in ecological studies. Mol Ecol. 12(1):251-62.
· Nubel U., Engelen B., Felske A., Snaidr J., Wieshuber A., Amann R.I., Wolfgang L. and Backhaus H. 1996. Sequence heterogeneities of genes encoding 16S rRNAs in Paenibacillus polymyxa detected by temperature gradient gel electrophoresis. J. Bacteriol. 178(19), 5636-5643.
· Satokari R.M., Vaughan E.E., Akkermans A.D.L., Saarela M., and de Vos W.M. 2001. Bifidobacterial diversity in human faeces detected by genus specific PCR and denaturing gradient gel electrophoresis. Appl. Environ. Microbiol. 67(2),504-513.
· Crosby, L. D., and C. S. Criddle, 2003. Understanding systematic error in microbial community analysis techniques as a result of ribosomal RNA (rrn) operon copy number. BioTechniques. 34(4), 790-803.

NON-RIBOSOMAL RNA GENE ANALYSES FOR BACTERIAL COMMUNITY ANALYSIS

· Dahllof, Baillie & Kjelleberg. 2000. rpoB based microbial communityanalysis avoids limitations inherent in 16S rRNA gene intraspeciesheterogeneity. Appl. Env. Micro. 66:3376-3380.

PCR ISSUES (There’s no end to these…)

Chimeras / Heteroduplex formation
Chimeras are always a concern in PCR analyses. To check to see if your sequence might be a chimera, you can use the chimera_check tool at the ribosomal database project (at least for ribosomal RNA gene sequences). You can also cut your sequence in half and BLAST each half of the sequence to see if you get similar results. There is also a new program called Pintail which is even more sophisticated.

http://www.cf.ac.uk/biosi/research/biosoft/Pintail/pintail.html
http://geta.life.uiuc.edu/RDP/misc/check_help.html
http://rdp8.cme.msu.edu/cgis/chimera.cgi?su=SSU

· Wang, G. C.-Y., and Y. Wang. 1997. Frequency of formation of chimeric molecules as a consequence of PCR coamplification of 16S rRNA genes from mixed bacterial genomes. Appl. Environ. Microbiol. 63:4645–4650.

“…Here evidence is presented for heteroduplexes as a major source of artifacts in mixed-template PCR…Heteroduplexes became increasingly prevalent as primers became limiting and/or template diversity was increased. … the diversity of artifactual sequences increases exponentially with the number of both variable nucleotides and of original sequence variants. Our model illustrates how minimization of heteroduplex molecules before cloning may reduce artificial genetic diversity detected during sequence analysis by clone screening. Thus, we developed a method to eliminate heteroduplexes from mixed-template PCR products by subjecting them to ‘reconditioning PCR’, a low cycle number re-amplification of a 10-fold diluted mixed-template PCR product. This simple modification to the protocol may ensure that sequence richness encountered in clone libraries more closely reflects genetic diversity in the original sample.”

· Janelle R. Thompson, Luisa A. Marcelino and Martin F. Polz. 2002. Heteroduplexes in mixed-template amplifications: formation, consequence and elimination by ‘reconditioning PCR’. Nucleic Acids Research. 30(9): 2083-2088.

Degenerate primers
Some functional gene (i.e. non ribosomal RNA gene) primers are highly degenerate and this can cause problems during DGGE analyses. Since the whole point of DGGE is to separate fragments that differ in sequence, identical PCR fragments that have different primer sequences can sometimes generate multiple bands on DGGE. The problem originates during the PCR reaction. When using degenerate primers, a low annealing temperature must be used to accommodate all the possible primer combinations (there is no point in having degenerate primers and then using an annealing temperature too high for some of the primers). However, at the low annealing temperature, some of the primers can anneal non-stringently to DNA target, and thus the non-stringent primer becomes incorporated into the growing DNA fragment. So, multiple primer combinations can anneal to the same template DNA and generate copies of the same fragment, but with different primer sequences. If these primer sequences are great enough, DGGE analysis will separate out the identical PCR fragments by the differences existing in the primer region. Sequencing/clone DGGE analysis can help resolve this issue. However, there isn’t much that can be done to avoid this problem – and it may again complicate measurements of diversity or dendogram analysis of DGGE gels. As a side note, this is the reason that you should NEVER submit the primer region of your sequence to Genbank.

Double bands
This appears to be another major complaint in the DGGE literature. Since many of the issues with double banding are a result of the PCR stage, you may want to consider:
>> Longer final elongation time (5-30 minutes of 72C)
>> Slow touchdown to 4C after final elongation stage
>> Check primer degeneracy
>> Concentration of DNA added to reaction
>> number of PCR cycles.

· Janse I, Bok J, Zwart G .2004. A simple remedy against artifactual double bands in denaturing gradient gel electrophoresis. Journal of microbiological methods. 57:279-281.

PCR Inhibition due to humics and polysaccharides
DGGE analyses can often be limited by the PCR step of the process. The PCR step can, in turn, be limited by the quality of the DNA extraction. Environmental samples rich in humic acids (organic rich soils, composts, decaying litter, etc.) and polysaccharides (biofilms, cyanobacteria, microbial mats, etc.) can contribute to poor quality DNA extracts. A lot of effort has been expended to deal with such environmental contaminants. In general, phenol/chloroform extractions and ultracentrifugation in a Cesium chloride gradient are the most effective for recovering pure DNA, these methods can be tiresome and contain toxic chemicals.

For humic acids, MoBio has a very nice kit for extracting soil DNA: The MoBio “PowerSoil” DNA isolation kit. I have also noticed that you can reduce the amount of humic acids that you recover in your extracts by removing EDTA from the extraction buffer, and by repeated cleaning of the DNA with guanidine thiocyanate, or by cleanup on PVPP columns. For a reference, see the following article; I would be happy to provide a more detailed protocol if necessary. I have read that “T4Gene” 32 protein can be added to PCR reactions to reduce humic inhibition.
· Inbar, E., Green, S.J., Hadar, Y. and D. Minz. 2005. Competing Factors of Compost Concentration and Proximity to Root Affect the Distribution of Streptomycetes. Microbial Ecology. 50:73-81.
· LaMontagne MG, Michel FC Jr, Holden PA, Reddy CA. 2002. Evaluation of extraction and purification methods for obtaining PCR-amplifiable DNA from compost for microbial community analysis. J Microbiol Methods. 2002 May;49(3):255-64.
· CC Tebbe and W Vahjen. 1993. Interference of humic acids and DNA extracted directly from soil in detection and transformation of recombinant DNA from bacteria and a yeast. Appl. Environ. Microbiol., 59(8): 2657-2665.

For polysaccharides, I recommend a potassium ethyl xanthogenate method.
· Tillett, D. and Neilan, B.A. 2000. Xanthogenate nucleic acid isolation from cultured and environmental cyanobacteria. J. Phycol. 36:251-258.

You can also try to overcome PCR inhibition due to contaminants by application of a “pre-PCR” stage using a specialized polymerase to make a large number of copies of genomic DNA.

· Gonzalez et al. 2005. Multiple displacement amplification as a pre-polymerase chain reaction (pre-PCR) to process difficult to amplify samples and low copy number sequences from natural environments. Environmental Microbiology. 7(7):1024-1028.

PCR Contamination
PCR contamination is a common event in molecular labs. This is particularly true for general bacterial PCR primers which will amplify any bacterial DNA that could be floating around in your lab. The first step to dealing with such contamination is to throw away all your PCR reagents (if this isn't too painful). I tend to use aliquots of each reagent and after having opened a tube I either dispose of the tube or use it for a less contamination likely PCR reaction (i.e. with a specific gene primer). You should think about where the contamination could come from. If one of your stock solutions (say primer stock) has been contaminated, you're in trouble. You can try a little test in somebody else's lab and PCR each of your reagents. You may also have some sort of contamination on your pipettes. Some researchers also use filters to clean up contamination - this is a last resort effort. To do this, you can filter all your "master mix" for PCR (WITHOUT THE ENZYME) through a 30 KD filter. The filter will pass the PCR primers, but will not pass the polymerase or genomic DNA (thus, add the enzyme afterwards). You can also work with filter tips that are not autoclaved but are ordered DNA/RNAse free; likewise with PCR tubes. Use purchased DNAse free PCR water. If none of that works, find a new career....

· Meier et al. 1993. Elimination of Contaminating DNA within Polymerase Chain Reaction Reagents: Implications for a General Approach to Detection of Uncultured Pathogens. Journal of Clinical Microbiology, 31:646-652.

Primer cleanup
When ordering primers I have never used anything but the most standard cleaning (i.e. desalting) offered by the companies. I have never found any improvement in DGGE analyses with reverse-phase HPLC, PAGE, or reverse-phase cartridge (RP1). Others may have different experience. Dr. A.G.C.L. (Arjen) Speksnijder reports that they have had poor experience with HPLC cleanup.

Primer Mistmatches / PCR Bias

· Kousuke Ishii and Manabu Fukui. 2001. Optimization of Annealing Temperature To Reduce Bias Caused by a Primer Mismatch in Multitemplate PCR. Appl. Environ. Microbiol. 67(8): 3753–3755.
· Shinya Kurata, Takahiro Kanagawa, Yukio Magariyama, Kyoko Takatsu, Kazutaka Yamada, Toyokazu Yokomaku, and Yoichi Kamagata. 2004. Reevaluation and Reduction of a PCR Bias Caused by Reannealing of Templates. Appl. Environ. Microbiol. 70(12):7545-9.


Primer Design
· Melt Profile Program: http://web.mit.edu/osp/www/melt.html
· Primernet (http://www.primernet.com/)
· Primrose : KE Ashelford, AJ Weightman & JC Fry. 2002. PRIMROSE: a computer program for generating and estimating the phylogenetic range of 16S rRNA oligonucleotide probes and primers in conjunction with the RDP-II database. Nucleic Acids Research, Vol. 30, No. 15 3481-3489
o http://www.cardiff.ac.uk/biosi/research/biosoft/Primrose/primrose.html
· Probe Library (ARB): http://www.arb-home.de/

Quantification
· Park JW, Crowley DE. 2005. Normalization of soil DNA extraction for accurate quantification of target genes by real-time PCR and DGGE. Biotechniques. 38(4):579-86.
· Bruggemann J, Stephen JR, Chang YJ, Macnaughton SJ, Kowalchuk GA, Kline E, White DC. 2000. Competitive PCR-DGGE analysis of bacterial mixtures: an internal standard and an appraisal of template enumeration accuracy. J Microbiol Methods. 40(2):111-23.
· D.G. Petersen and I. Dahllöf. 2005. Improvements for comparative analysis of changes in diversity of microbial communities using internal standards in PCR-DGGE. FEMS Ecology. [In Press].

Optimization
· Wood GS, Uluer AZ.1999. Polymerase chain reaction/denaturing gradient gel electrophoresis (PCR/DGGE): sensitivity, band pattern analysis, and methodologic optimization. Am J Dermatopathol. 21(6):547-51.
· Markus M. Moeseneder, Jesús M. Arrieta, Gerard Muyzer, Christian Winter, and Gerhard J. Herndl. 1999. Optimization of Terminal-Restriction Fragment Length Polymorphism Analysis for Complex Marine Bacterioplankton Communities and Comparison with Denaturing Gradient Gel Electrophoresis. Applied and Environmental Microbiology, 65(8): 3518-3525.
· Vanessa M. Hayes, Ying Wu, Jan Osinga, Inge M. Mulder, Pieter van der Vlies, Peter Elfferich, Charles H. C. M. Buys, Robert M. W. Hofstra. 1999. Improvements in gel composition and electrophoretic conditions for broad-range mutation analysis by denaturing gradient gel electrophoresis. Nucleic Acids Research. 27(20): 29.

General PCR articles
· Speksnijder AG. 2001. Microvariation artifacts introduced by PCR and cloning of closely related 16S rRNA gene sequences Appl Environ Microbiol. 67(1):469-72

PERPENDICULAR GELS
Optimization of the gradient for DGGE analyses can be done empirically by running an initial gel with a 0-80% gradient and then narrowing the gradient in subsequent analyses by inspection of the first gel. Alternatively, you can pour a perpendicular gel. In this case, during the casting stage the gel is turned on its side. After the gel has polymerized, the gel is set upright, and a single sample is analyzed. This can help identify which gradient, or if any gradient, will separate the fragments of interest. Protocols for pouring these gels come with the Bio-Rad system, but I don’t think the Ingeny system comes with the capacity to pour such a gel.

POLYMERIZATION
Troubleshooting DGGE gels that do not look good can include problems with the pouring and polymerization of your acrylamide. In addition to checking that your reagents are fresh, you may want to look at the following website:
· http://www.bio-rad.com/LifeScience/pdf/Bulletin_1156.pdf

You may also want to add commercially available substances to your acrylamide to enhance the strength of your gel and reduce tearing. Some users have complained that this, or silanizing glass plates made DGGEs worse. Oscar J. de Vos from Wageningen University indicates that UV light cannot pass through the Gelbond product listed below. Still, you may want to check this out:
http://www.cambrex.com/Content/bioscience/CatNav.oid.520.prodoid.GelbondPAG

There has been some discussion in the DGGE group about how long you should let your gels polymerize. In general, 1.5 hr seems to be the absolute minimum. You should also not move your gel during polymerization, if possible. Some users have suggested that 2 hr polymerization is adequate and should not be longer than this. I have not found that length of polymerization time before running is particularly important, provided the 1.5 hr time is reached. If storing gels, I wouldn’t recommend more than 1 or 2 days. We used to put the gels in a plastic bag with a moist kimwipe when storing overnight. I have left gels out of the refrigerator overnight without any detrimental effect. You should make sure that there is some moist towel or kimwipe to ensure that the gel does not dry out.

SEPARATION OF BANDS
If you are not getting good separation on your DGGE, there can be a number of issues to examine. First, not all genes and regions of genes are suited to DGGE analyses. There is a rule of thumb which suggests that if DNA fragments do not differ by 1% or more (is it true?), it will be difficult to resolve them by DGGE. You can reduce the size of the PCR fragment, and this may increase separation if you don’t eliminate variable sites. You should check, if possible, available sequences to see if there are reasonable differences in sequence that would allow separation. Also, remember that in general, large fragments (>600 bp) do not separate well by DGGE. There is the 1650 bp fragment of fungal 18S ribosomal RNA, but this appears to be a freakish exception.
· E.J. VAINIO and J. HANTULA. 2000. Direct analysis of wood-inhabiting fungi using denaturing gradient gel electrophoresis of amplified ribosomal DNA. Mycological Research 104: 927-936.
>> An improper gradient can yield poor DGGE results. If the gradient is too wide, band may migrate very closely and you will loose reasonable separation; however, these gels tend to have the sharpest bands and look very nice. One should be careful not to make too narrow a gradient as this can yield fuzzy bands.
>> Check to see that the voltage and run time are adequate.
>> Check to see that there are no blockages to current running through the system (check that the amperage is appropriate for your system).
>> Did you remember to put a GC clamp on one of the primers?
>> Did you put a GC clamp on both primers? If so, whoops…
>> Do you have more than one sequence in your system? If not, perhaps you should be getting a single band…

· Y Wu, VM Hayes, J Osinga, IM Mulder, MW Looman, CH Buys, and RM Hofstra. 1998. Improvement of fragment and primer selection for mutation detection by denaturing gradient gel electrophoresis. Nucleic Acids Research. 26:5432-5440.
· Kisand V, Wikner J. 2003. Limited resolution of 16S rDNA DGGE caused by melting properties and closely related DNA sequences. J Microbiol Methods. 54(2):183-91.

SINGLE STRANDED DNA
Single stranded DNA can sometimes cause problems with DGGE. If this is your particular problem, you can try digesting it away prior to loading your PCR product on DGGE using Mung Bean nuclease.

SMILING GELS
Smiling of bands near the edges of DGGE gels appears to be endemic in all systems. While the exact cause of this is not entirely clear, the smiling effect can be held in check by two approaches, best used together:
· Don’t load PCR product in the very far lanes.
· Apply grease to the spacers. Older spacers may require more grease than new spacers, but don’t overload the spacers with grease. A thin film is adequate.

· Brinkhoff, T. and Van Hannen, E.J. 2001. Use of Silicone Grease to Avoid 'smiling Effect' in DGGE. Journal of Rapid Methods and Automation in Microbiology 9:259-261.

STORING GELS AFTER ELECTROPHORESIS
According to James Hollibaugh, DGGE gels can be stored for subsequent extraction of DNA by drying them onto filter paper using a gel dryer. Once dry they can be stored at room temperature in a loose-leaf binder (use sheet protectors) for at least a year.
· Hollibaugh, J. T., P. S. Wong, N. Bano, S. K. Pak, E. M. Prager and C. Orrego. 2001. Stratification of microbial assembledges in Mono Lake, California, and response to a mixing event. Hydrobiologia 466:45-60.

WELLS – WAVY, POOR QUALITY, ETC.
Sometimes the wells that are formed when you remove the comb are of poor quality, usually because the polymerized acrylamide that forms the walls between the wells does not stand up straight and collapses. You can rectify this after the fact by using a syringe tip or other similar device to manually straighten each well.

To avoid this problem in general, I have found that one of the causes of this is to pour TOO MUCH acrylamide into the comb area (meaning that you over-pour the amount of acrylamide in the stacking gel area). For whatever reason, if you are careful to add just the amount needed, you’ll have less problems. Remember not to add too little, because acrylamide shrinks as it polymerizes. Putting plastic wrap over the top of the gel has been suggested to reduce evaporation and limit shrinkage.

Also, pull your comb out very slowly.

DGGE OF HIGH GC PCR FRAGMENTS
· Per Guldberg1, Kirsten Grønbæk, Anni Aggerholm, Anton Platz, Per thor Straten, Vibeke Ahrenkiel, Peter Hokland and Jesper Zeuthen. 1998. Detection of mutations in GC-rich DNA by bisulphite denaturing gradient gel electrophoresis. Nucleic Acids Research. 26(6): 1548-1549.
· Ying Wu, Rein P. Stulp, Peter Elfferich, Jan Osinga, Charles H. C. M. Buys, Robert M. W. Hofstra. 1999. Improved mutation detection in GC-rich DNA fragments by combined DGGE and CDGE. . Nucleic Acids Research. 27(15): 9.


FANCY DGGE TECHNIQUES
· Van Orsouw NJ, Vijg J. 1999. Design and application of 2-D DGGE-based gene mutational scanning tests. Genet Anal. 14(5-6):205-13.
· Nathalie J. van Orsouw, Rahul K. Dhanda, R. David Rines, Wendy M. Smith1, Iakovos Sigalas, Charis Eng1, Jan Vijg. 1998. Rapid design of denaturing gradient-based two-dimensional electrophoretic gene mutational scanning tests. Nucleic Acids Research. 26(10): 2398-2406.
· Green, S.J. and D. Minz. 2005. Suicide Polymerase Endonuclease Restriction (SuPER) – a novel technique for enhancing PCR amplification of minor DNA templates. Appl. Environ. Microbiol. 71:4721-4727.
· Cremonesi L, Firpo S, Ferrari M, Righetti PG, Gelfi C. 1997. Double-gradient DGGE for optimized detection of DNA point mutations. Biotechniques. 22(2):326-30.
· Burmeister, M., diSibio, G., Cox, D. R., and Myers, R. M. Identification of polymorphisms by genomic denaturing gradient gel electrophoresis: application to the proximal region of human chromosome 21. Nucleic Acids Res 19(7): 1475–81, 1991.
· William E. Holben, Kevin P. Feris, Anu Kettunen, and Juha H. A. Apajalahti. 2004. GC Fractionation Enhances Microbial Community Diversity Assessment and Detection of Minority Populations of Bacteria by Denaturing Gradient Gel Electrophoresis. Appl Environ Microbiol. 70(4): 2263–2270.

RECOMMENDED PCR-DGGE PRIMER SETS AND GRADIENT CONDITIONS

Bacterial 16S rRNA gene: 341F / 907R (E. coli numbering. See Muyzer References; Primer in bold):

341F-GC: 5’-CGC CCG CCG CGC CCC GCG CCC GTC CCG CCG CCC CCG CCC GCC TAC GGG AGG CAG CAG-3’
907R: CCG TCA ATT CMT TTG AGT TT

PCR conditions: 4.0 mM Mg; 60C Annealing temperature
Nest-PCR conditions: 4.0 mM Mg; 64C Annealing temperature; 28 cycles
DGGE conditions: 25/30 to 60/70% denaturant; 100V; 60C; 17 hr; 6% acrylamide


Keywords: denaturing, gradient, gel electrophoresis, help, advice, guide, DGGE, community composition, molecular biology, fingerprinting, PCR-DGGE, nested PCR.

133 Comments:

At 3:57 PM, Blogger jsfoster said...

Dude, i can't wait to read your RTPCR web site when you get that up and running ;-)

It looks awesome. I will have to link your site to my web page when ever I get mine done.

toodles
Jamie

 
At 11:25 PM, Anonymous Raj said...

I must say! tremendous amounts of effort put in explaining DGGE... Great work... I am sure many will use this as a guide. You will be famous one day.

 
At 1:14 AM, Anonymous Anonymous said...

Thank you so much for a clear and thorough DGGE protocol!! :)

 
At 6:35 AM, Anonymous Anonymous said...

Absolutely brilliant!
:D

 
At 12:32 PM, Anonymous Anonymous said...

Fantastic blogg! I find it extremely useful, thanks so much for doing this!!

 
At 3:57 PM, Blogger Dexter said...

Thank you guys very much for building up this DGGE technology specific website. It's very great because I found a lot information need for my research!

 
At 7:20 AM, Blogger Rok said...

Good job! Well done! I will recommend your blog to my colleagues and students.

Rok

 
At 1:39 AM, Anonymous Anonymous said...

Not sure what you mean-

'..what is the point? I would like to suggest that a better way to approach this is to clone directly from the original PCR product (using GC-clamped PCR primers) and then screen the clones against the environmental sample...'

Can somebody elaborate?

 
At 10:59 AM, Anonymous Anonymous said...

Thank you very much.

This blog is fantastic. It was very useful for me.

Congratulations and I hope that you continue writting about this topics.

 
At 11:30 PM, Anonymous Anonymous said...

Hi Stefan,

I`d like to thank you first for the great jib!
I got a Q, At first I have no bubbles trapped but after a few hours having it run, bubbles appear and get stuck in the middle of the gel!!! What`s wrong?

I appreciate it in advance.
(Plz email me your reply if possible at sam_phd2006@yahoo.com)


Cheers,

Sam

 
At 8:50 PM, Blogger someone said...

i have a question about DGGE. can anyone answer me , is formamide neccessary for DGGE?

 
At 8:14 PM, Anonymous Anonymous said...

does anyone know how to best clean plates after using silicone grease on the spacers? is there a particular solvent that works for this? so far i've only succeeded in smearing the stuff all over the plates.

 
At 3:37 AM, Anonymous Anonymous said...

A very good guide. Thanks.

 
At 10:10 AM, Anonymous Anonymous said...

Is there anyone that can help me with post DGGE? I've excised the bands from the DGGE gel, and washed with ddwater and then eluted overnight with elution buffer. I then used the top layer as template DNA in a PCR with the same DGGE primers but without the GC clamp. Im getting double bands on the agarose gel. wat could be the reason for this and how can I eliminate it?

Thanks alot!

 
At 1:00 PM, Blogger Stefan J. Green said...

In response to the most recent comment - I would suggest that you try to raise the annealing temperature of your post-DGGE amplification, or dilute the DNA template (try a range of dilutions), or decrease the number of PCR cycle. You can often get spurious bands when working with very high concentrations of DNA.
Let me know if that helps,
Stefan

 
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At 12:01 PM, Anonymous Anonymous said...

Nice work and it is VERY helpful! I will recommend it to my colleagues.

 
At 7:42 PM, Anonymous Panic Attack said...

This DGGE technology specific is a great aid to research. It must have took a lot of time and effort on your part to deliver to us such a detailed and comprehensive report. I appreciate the clarity of presentation of the DGGE protocol.

 
At 2:49 AM, Anonymous ANU said...

VERY INFORMATIVE AND THANKS .......EXPLANATION IS VERY CLEAR...NICE TIPS TOO...

 
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So, I don't actually believe this will work.

 
At 4:33 AM, Anonymous Rahul said...

Hii Stefan,
Thanks for the guide its really helpful. I have just started with DGGE and was finding it tough to optimize untill I read this. Currently I am able to amplify community DNA with GC clamps and make a decent gradient gel as well. The only recurring problem is I am not to make proper wells (lack of polymerization due to oxygen presence I guess). I use 0.09% (25 ul) each of TEMED and 10% APS for a 20 ml gel as directed in BioRad manual. Have tried with higher volumes as well. 100 ul APS and 50 ul TEMED doesnt give me sufficient time to cast. Please help me save precious resources and time in optimization.

Thanks.

 
At 11:42 AM, Anonymous Anonymous said...

Dear Stefan,
If, I should compare different soils, (comparing different fingerprinting or asses the different diversity index) how much DNA I should add to pcr reaction? Should I measure, with the spectrophotometer, the dna amount of each sample and add the same amount in each pcr? Or Should I take the same amount of soil (5 g for instance) perform the dna extraction and purification and add directly 2 (for instance) microliter of dna in each pcr?"

 
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At 9:57 PM, Anonymous Anonymous said...

I am using Ingeny for DGGE. Is it advisable to run DGGE for 2 specific primer (with different acrylamide gel percentage) at the same time (front and back gel)? Thank you. I wish to excise the band.

 
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