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DNA Sequencing and Gene Analysis Determining DNA Sequence • Originally 2 methods were invented around 1976, but only one is widely used: invented by Fred Sanger. – After discussing Sanger sequencing, we will go over the newer pyrosequencing method. • Uses DNA polymerase to synthesize a second DNA strand that is labeled. DNA polymerase always adds new bases to the 3’ end of a primer that is base-paired to the template DNA. • Also uses chain terminator nucleotides: dideoxy nucleotides (ddNTPs), which lack the -OH group on the 3' carbon of the deoxyribose. When DNA polymerase inserts one of these ddNTPs into the growing DNA chain, the chain terminates, as nothing can be added to its 3' end. Sanger Sequencing Reaction • The template DNA is usually single stranded DNA, which can be produced from plasmid cloning vectors that contain the origin of replication from a single stranded bacteriophage such as M13 or fd. The primer is complementary to the region in the vector adjacent to the multiple cloning site. • Sequencing is done by having 4 separate reactions, one for each DNA base. All 4 reactions contain the 4 normal dNTPs, but each reaction also contains one of the ddNTPs. In each reaction, DNA polymerase starts creating the second strand beginning at the primer. When DNA polymerase reaches a base for which some ddNTP is present, the chain will either: – terminate if a ddNTP is added, or: – continue if the corresponding dNTP is added. – which one happens is random, based on ratio of dNTP to ddNTP in the tube. However, all the second strands in, say, the A tube will end at some A base: you get a collection of DNAs that end at each of the A's in the region being sequenced. • • • • Electrophoresis • • • The newly synthesized DNA from the 4 reactions is then run (in separate lanes) on an electrophoresis gel. The DNA bands fall into a ladderlike sequence, spaced one base apart. The actual sequence can be read from the bottom of the gel up. Automated sequencers use 4 different fluorescent dyes as tags attached to the dideoxy nucleotides and run all 4 reactions in the same lane of the gel. – Today’s sequencers use capillary electrophoresis instead of slab gels. – Radioactive nucleotides (32P) are used for non-automated sequencing. • Sequencing reactions usually produce about 500-1000 bp of good sequence. Pyrosequencing • Recently a number of faster and cheaper sequencing methods have been developed. – The Archon X prize (2006): "the first Team that can build a device and use it to sequence 100 human genomes within 10 days or less, with an accuracy of no more than one error in every 100,000 bases sequenced, with sequences accurately covering at least 98% of the genome, and at a recurring cost of no more than $10,000 (US) per genome.” – Probably the most widely used new methods involve the pyrosequencing biochemical reactions (invented by Nyren and Ronaghi in 1996), with the massively parallel microfluidics technology invented by the 454 Life Sciences company. We can call this combined technology “454 sequencing”. • Applications: – – – – – sequencing of whole bacterial genomes in a single run sequencing genomes of individuals metagenomics: sequencing DNA extracted from environmental samples looking for rare variants in a single amplified region, in tumors or viral infections transcriptome sequencing: total cellular mRNA converted to cDNA. Pyrosequencing Biochemistry • • In DNA synthesis, a dNTP is attached to the 3’ end of the growing DNA strand. The two phosphates on the end are released as pyrophosphate (PPi). ATP sulfurylase uses PPi and adenosine 5’phosphosulfate to make ATP. – • ATP sulfurylase is normally used in sulfur assimilation: it converts ATP to adenosine 5’phosphosulfate and PPi. However, the reaction is reversed in pyrosequencing. Luciferase is the enzyme that causes fireflies to glow. It uses luciferin and ATP as substrates, converting luciferin to oxyluciferin and releasing visible light. – The amount of light released is proportional to the number of nucleotides added to the new DNA strand. • After the reaction has completed, apyrase is added to destroy any leftover dNTPs. More Pyrosequencing • The four dNTPs are added one at a time, with apyrase degradation and washing in between. • The amount of light released is proportional to the number of bases added. Thus, if the sequence has 2 A’s in a row, both get added and twice as much light is released as would have happened with only 1 A. • The pyrosequencing machine cycles between the 4 dNTPs many times, building up the complete sequence. About 300 bp of sequence is possible (as compared to 800-1000 bp with Sanger sequencing). • The light is detected with a charge-coupled device (CCD) camera, similar to those used in astronomy. • YouTube animation (with music!): http://www.youtube.com/watch?v=kYAGFrbGl6E 454 Technology • To start, the DNA is sheared into 300-800 bp fragments, and the ends are “polished” by removing any unpaired bases at the ends. • Adapters are added to each end. The DNA is made single stranded at this point. • One adapter contains biotin, which binds to a streptavidin-coated bead. The ratio of beads to DNA molecules is controlled so that most beads get only a single DNA attached to them. • Oil is added to the beads and an emulsion is created. PCR is then performed, with each aqueous droplet forming its own micro-reactor. Each bead ends up coated with about a million identical copies of the original DNA. More 454 Technology • After the emulsion PCR has been performed, the oil is removed, and the beads are put into a “picotiter” plate. Each well is just big enough to hold a single bead. • The pyrosequencing enzymes are attached to much smaller beads, which are then added to each well. • The plate is then repeatedly washed with the each of the four dNTPs, plus other necessary reagents, in a repeating cycle. • The plate is coupled to a fiber optic chip. A CCD camera records the light flashes from each well. Illumina/Solexa Sequencing • Another high throughput Next Generation Sequencing method. • http://www.youtube.com/watch?v=77r5p8IBwJk&NR=1 • http://www.youtube.com/watch?v=HtuUFUnYB9Y • First we will discuss the sequencing reaction itself, and then deal with how it is done on many sequences simultaneously. Illumina Sequencing Chemistry • • this method uses the basic Sanger idea of “sequencing by synthesis” of the second strand of a DNA molecule. Starting with a primer, new bases are added one at a time, with fluorescent tags used to determine which base was added. The fluorescent tags block the 3’-OH of the new nucleotide, and so the next base can only be added when the tag is removed. – • So, unlike pyrosequencing, you never have to worry about how many adjacent bases of the same type are present. The cycle is repeated 50-100 times. Illumina Massively Parallel System • The idea is to put 2 different adapters on each end of the DNA, then bind it to a slide coated with the complementary sequences for each primer. This allows “bridge PCR”, producing a small spot of amplified DNA on the slide. • The slide contains millions of individual DNA spots. The spots are visualized during the sequencing run, using the fluorescence of the nucleotide being added. Sequence Assembly • • DNA is sequenced in very small fragments: at most, 1000 bp. Compare this to the size of the human genome: 3,000,000,000 bp. How to get the complete sequence? In the early days (1980’s), genome sequencing was done by chromosome walking (aka primer walking): sequence a region, then make primers from the ends to extend the sequence. Repeat until the target gene was reached. – The cystic fibrosis gene was identified by walking about 500 kbp from a closely linked genetic marker, a process that took a long time and was very expensive. – Still useful for fairly short DNA molecules, say 1-10 kbp. Shotgun Sequencing • • Shotgun sequencing is what is typically done today: DNA is fragmented randomly and enough fragments are sequenced so each base is read 10 times or more on average. The overlapping fragments (“reads”) are then assembled into a complete sequence. For large genomes, hierarchical shotgun sequencing is a useful technique: first break up the genome into an ordered set of cloned fragments (scaffolds), usually BAC clones. Each BAC is shotgun sequenced separately. Assembly Problems • • • • • • In principle, assembling a sequence is just a matter of finding overlaps and combining them. In practice: – most genomes contain multiple copies of many sequences, – there are random mutations (either naturally occurring cell-to-cell variation or generated by PCR or cloning), – there are sequencing errors and misreadings, – sometimes the cloning vector itself is sequenced – sometimes miscellaneous junk DNA gets sequenced Getting rid of vector sequences is easy once you recognize the problem: just check for them. Repeat sequence DNA is very common in eukaryotes, and sequencing highly repeated regions (such as centromeres) remains difficult even now. High quality sequencing helps a lot: small variants can be reliably identified. Sequencing errors, bad data, random mutations, etc. were originally dealt with by hand alignment and human judgment. However, this became impractical when dealing with the Human Genome Project. This led to the development of automated methods. The most useful was the phred/phrap programs developed by Phil Green and collaborators at Washington University in St. Louis. Phred Quality Scores • Phred is a program that assigns a quality score to each base in a sequence. These scores can then be used to trim bad data from the ends, and to determine how good an overlap actually is. – there are much improved algorithms now, but the phred quality score is still widely used. • Phred scores are logarithmically related to the probability of an error: a score of 10 means a 10% error probability; 20 means a 1% chance, 30 means a 0.1% chance, etc. – Q = -10 log P, where Q is the phred score and P is the probability that the base was called incorrectly. – A score of 20 is generally considered the minimum acceptable score. • Phred uses Fourier analysis (decomposing the data into a series of sine waves) to examine chromatogram trace data. – – – • • First, find the expected position of each peak, assuming they are supposed to be evenly spaced, with no compressions or other factors altering peak positions. Next, find actual center and area of each peak. Finally, match observed peaks to expected. This involves splitting some peaks and ignoring others. Assigning a quality score involves comparing various parameters determined for each peak with data that was generated from known sequences run under a wide variety of conditions (i.e., based on ad hoc observations and not theory). Since all four traces (A, C, G, T) are examined separately, phred generates a best-guess sequence. Phred output is a file where each line contains one base and its quality score. Phred Algorithm Combining Sequences with Phrap • • • • • Phrap first examines all reads for matching “words”: short sections of identical sequence. The matching words need to be in the same order and spacing. – Sequences in both orientations are examined, using the reverse-complement sequence if necessary. The entire sequences of pairs with matching words are then aligned using the SmithWaterman algorithm (a standard technique we will look at later). Phrap then looks for discrepancies in the combined sequences, using phred scores to decide between alternatives. Phrap generates quality scores from the combined phred data. – Sequencing errors are not necessarily random: homopolymeric regions (several of the same base in a row) are notoriously tricky to sequence accurately. Using the opposite strand often helps resolve these regions. Also using a different sequencing technology or chemistry. Sequences are combined with a greedy algorithm: all pairs of fragments are scored for the length and quality of their overlap region, and then the largest and bestmatched pair is merged. This process is repeated until some minimum score is reached. The result is a set of contigs: reads assembled into a continuous DNA sequence. – The ideal result is the entire chromosome assembled into a single contig. Finishing the Sequence • Shotgun sequencing of random DNA fragments necessarily misses some regions altogether. – Also, for sequencing methods that involve cloning (Sanger), certain regions are impossible to clone: they kill the host bacteria. • • Thus it is necessary to close gaps between contigs, and to re-sequence areas with low quality scores. This process is called finishing. It can take up to 1/2 of all the effort involved in a genome sequencing project. Mostly hand work: identify the bad areas and sequence them by primer walking. – Sometimes using alternative sequencing chemistries (enzymes, dyes, terminators, dNTPs) can resolve a problematic region. • Once a sequence is completed, it is usually analyzed by finding the genes and other features on it: annotation. • Submission of the annotated sequence to Genbank allows everyone access to it: the final step in the scientific method. Single Nucleotide Polymorphisms • • • • Looking at many individuals, you can see that most bases in their DNA are the same in everyone. However, some bases are different in different individuals. These changes are single nucleotide polymorphisms (SNPs). SNPs are found everywhere in the genome, and they are inherited in a regular Mendelian fashion. These characteristics makes them good markers for finding disease genes and determining their inheritance. Lots of ways to detect SNPs, many of which are easy to automate. Primer extension: make a primer 1 base short of the SNP site, and then extend the primer using DNA polymerase with nucleotides having different fluorescent tags. Gene Detection • It is surprisingly hard to be sure that a given genomic sequence is a gene: that it is ever expressed as RNA. • Protein-coding regions are open reading frames (ORFs): they don’t contain stop codons. • But, human genes often contain long introns and very short exons, and some parts of genes are introns in one cell type but exons in other cell types. So, finding all the pieces of a gene can be a challenge. • Three questions: – is a given DNA sequence ever expressed? – is the sequence expressed in a given cell type or set of conditions? – what is the intron/exon structure of the sequence? Evolutionarily Conserved Sequences • • • • When looking across different species, most DNA sequences are not conserved. However, the exons of genes are often highly conserved, because their function is necessary for life. Zoo blot: a Southern blot containing genomic DNA from many species. Probe it with the sequence in question: exons will hybridize with other species’ DNA, while introns and non-gene DNA won’t. Computer-based homology search: BLAST search. Do similar sequences appear in the nucleotide databases? Especially chimpanzee and mouse, which have complete genome sequences available. Detecting Gene Expression • Northern blots: RNA extracted from various tissues or experimental conditions, run on an electrophoresis gel, then probed with a specific DNA sequence. Detecting Gene Expression • Real time PCR: – first convert all mRNA in a sample to cDNA using reverse transcriptase, – then amplify the region of interest using specific primers. – Use a fluorescent probe to detect and quantitate the specific product as it is being made by the PCR reaction. – the two components of the fluorescent tag interact to quench each other. When one part is removed by the Taq polymerase, the quenching stops and fluorescence can be detected. Expressed Sequence Tags • ESTs are cDNA clones that have has a single round of sequencing done from one end. • First extract mRNA from a given tissue. Then convert it to cDNA and clone. • Sequence thousands of EST clones and save the results in a database. • A search can then show whether your sequence was expressed in that tissue. – quantitation issues: some mRNAs are present in much higher concentration than others. Many EST libraries are “normalized” by removing duplicate sequences. • Also can get data on transcription start sites and exon/intron boundaries by comparing to genomic DNA – but sometimes need to obtain the clone and sequence the rest of it yourself. RNA Seq • • This is a new method, published in 2008. It is probably the method of choice today for analyzing RNA content. Also called whole transcriptome shotgun sequencing. Very simple: isolate messenger RNA, break it into 200-300 base fragments, reverse transcribe, then perform large scale sequencing using 454, Illumina. Or other massively parallel sequencing technology. – RNA sequences then compared to genomic sequences to find which gene is expressed and also exon boundaries – Exon boundaries are a problem with very short reads: you might only have a few bases of overlap to one of the exons. • • As with all RNA methods, which RNAs are present depends on the tissue analyzed and external conditions like environmental stress or disease state. Get info on copy number over a much wider range than microarrays. Also detects SNPs. Etc. • New techniques in DNA/RNA technology are being developed constantly. The main goal is to increase reliability and decrease cost. Primarily the aim is to automate as much as possible. • Just a few techniques we are not going to discuss: RACE, SAGE, differential display, S1 nuclease protection Protein Methods • • It is important to be sure that the protein product of a gene is made, and to know where in the tissue or cell it is made, and how much is made. Most protein detection is based on either making antibodies to the protein of interest, or by making a fusion protein: your protein fused to a fluorescent protein. – GFP: green fluorescent protein. Isolated from jellyfish. Several variants give different colors. It still works when it is fused to other proteins. • Often done in conjunction with confocal microscopy: examining the same image with visible light and fluorescence. Two-Dimensional Gel Electrophoresis • 2D gels are a way of separating proteins into individual spots that can be individually analyzed. – Proteins are first separated by their isoelectric point and then by their molecular weight. – Individual protein spots can then be identified using mass spectrometry • Isoelectric focusing separates proteins by their isoelectric point, the pH at which the net surface charge is zero. – IEF uses a mixture of ampholytes, chemical compounds that contain both acidic and basic groups. – When an electric field is applied, the ampholytes move to a position in the gel where their net charge is zero, and in the process they set up a pH gradient. – Proteins also move down the pH gradient until they reach a pH where they have no net charge, their isoelectric point. – Isoelectric focusing is thus an equilibrium process: proteins move to a stable position and stay there. (But in practical terms, the gradient breaks down over time). SDS-PAGE • SDS-PAGE is a method for separating proteins according to their molecular weight. – SDS = sodium dodecyl sulfate (a.k.a. sodium lauryl sulfate), a detergent that unfolds proteins and coats them in charged molecules so that their charge to mass ratio is essentially identical. • “Native” gel electrophoresis uses undenatured proteins, which vary greatly in charge to mass ratio. – SDS denaturation isn’t perfect: some proteins behave anomalously, – PAGE = polyacrylamide gel electrophoresis 2D gels • • • First, isoelectric focusing is performed on a protein sample, running the proteins through a narrow tube or strip of acrylamide. Then the IEF gel is placed on top of an SDS gel, allowing the proteins to be separated by their molecular weight at right angles to the isoelectric point separation. Then the gel is stained with a general protein stain such as Coomassie Blue or silver stain. – • A Western blot involves transferring the separated proteins onto a membrane, where specific proteins can be identified by antibody binding. A couple of issues: – – – – While a cell might contain up to 100,000 proteins, at best only 3000 spots can be resolved. Proteins expressed at a low level (such as regulatory proteins) don’t show up well: spot size is proportional to the amount present in the cell Special techniques are needed for membrane proteins, which aren’t easily solubilized by the usual techniques. Comparing spots between 2D gels require image analysis software (and well-standardized running conditions). Mass Spectrometry • The general principle of mass spectrometry is that if you ionize a group of atoms or molecules, you can separate them on the basis of charge to mass ratio, by accelerating them in an electric field in a vacuum. – The original mass spectrometers were used to separate isotopes, based on slightly different masses. • • During the ionization process, proteins tend to break up in characteristic ways, producing “molecular ions” whose molecular weights can be measured very precisely. Since you are generally working with an already sequenced genome, you can predict the size of fragments that will be generated by any gene. Thus you can identify the gene product by matching the actual fragments with list of predicted fragments. • For most protein work, the proteins are first digested into small fragments (say , 5-10 amino acids), separated by HPLC (high performance liquid chromatography), and then run individually through the mass spec. – – • • • Protein sequencing and older protein identification methods also start with proteolytic digestion Endopeptidases that digest at known sites are used, such as trypsin (cleaves after Lys or Arg) and chymotrypsin (cleaves after Phe, Trp, or Tyr). Ionizing the peptide needs to be done rather gently. One common technique is MALDI (matrixassisted laser desorption/ionization). The proteins are mixed with the matrix molecules, which efficiently absorb the UV laser energy and encourage ionization of the proteins. When irradiated with the laser, they vaporize along with the protein, but their small size makes them easy to detect and ignore. Time-of-flight mass spectrometry is generally used (so the whole thing is MALDI-TOF). The moelcular ions are accelerated in an electric field, and the time it takes them to cross a chamber of known length is proportional to their mass 9actaully, charge to mass ratio). This technique works well for the wide range of sizes seen with peptides. Sample comparisons can be done by labeling one sample with a heavy, stable isotope such as 13C or 15N. The samples are mixed before 2D electrophoresis and they co-migrate on the gel. However, mass spec can easily resolve them. More Mass Spectrometry Antibodies • • • • • If you inject rabbits (usually) with your protein, the rabbit will develop an immune response against it. This means that a set of immunoglobulin (Ig) molecules that specifically bind to your protein will be produced. Mostly IgG, the main immunoglobulin that circulates in the blood. Your protein is acting as an antigen. Each Ig molecule binds to a specific epitope on the protein. The blood serum from the rabbit contains polyclonal antibodies: several different Ig proteins that bind to different epitopes on your protein. Monoclonal antibodies can be made in mice. Each monoclonal antibody is a single IgG molecule, and so it will bind to a single epitope on your protein. Monoclonal antibodies are made by fusing individual Ig-producing spleen cells to myeloma cells, which creates an immortal Ig-producing cell line, called a hybridoma. ------------------------------------------------------------------------------- Western Blots • E. M. Southern, inventor of the Southern blot, is a real person. However, his cool name has been hijacked to name the Northern blot (i.e. running RNA instead of DNA on the gel), and the Western blot (running protein on the gel). – Also called a “protein immunoblot”. • Proteins are separated by gel electrophoresis. – • • Can be done with denatured proteins (SDS gel), which separates proteins by molecular weight, or under nondenaturing conditions (no SDS), which separates proteins by their surface charge and size. The proteins on the gel are then blotted onto a nitrocellulose membrane. The specific protein of interest is detected using antibodies. – – – First antibody is made in a rabbit (usually) by injecting the specific protein of interest. It binds to the protein on the blot. The presence of the first antibody is detected by using a labeled second antibody that has been conjugated (covalently bound to) a fluorescent tag or to an easily detected enzyme. The second antibody is made by injecting rabbit immunoglobulins into a sheep.