Last time you purified your 16S PCR product, reacted it with specially prepared DNA, and transformed the product into an engineered cell strain. Eight independent colonies were selected from each of your plates and grown overnight in liquid culture. You will isolate and sequence DNA from each colony, then pool your results with all other groups studying that particular bird sample and construct a phylogenetic tree representing the bacterial composition in that sample.
Last time we talked about the features of the transformation strain needed to synthesize and select for the whole plasmid, namely Cre recombinase and a lacZ mutation, respectively. Now let's talk about those features relevant to extracting DNA. As you can gather from the linked manual (PDF), the StrataClone SoloPack cells have endA and recA mutations, both of which make cloning go more smoothly. First, endA1 limits the non-specific destruction of plasmid (and chromosomal) DNA normally carried out by the EndA enzyme, thus maximizing DNA recovery. (The cells also have additional deficiencies in restriction-based nucleases.) Second, recA1 makes the cells incapable of homologous recombination, which could otherwise cause undesirable intermingling between the plasmid and chromosomal DNA or among plasmids.
The procedure for DNA isolation at this scale is commonly termed "mini-prep," which distinguishes it from a “maxi-prep” that involves a larger volume of cells and additional steps of purification. The overall goal of each prep is the same--to separate the plasmid DNA from the chromosomal DNA and cellular debris, allowing the plasmid DNA to be studied further. In the traditional mini-prep protocol, the media is removed from the cells by centrifugation. The cells are resuspended in a solution that contains Tris to buffer the cells and EDTA to bind divalent cations in the lipid bilayer, thereby weakening the cell envelope. A solution of sodium hydroxide and sodium dodecyl sulfate (SDS) is then added. The base denatures the cell’s DNA, both chromosomal and plasmid, while the detergent dissolves the cellular proteins and lipids. The pH of the solution is returned to neutral by adding a mixture of acetic acid and potassium acetate. At neutral pH the SDS precipitates from solution, carrying with it the dissolved proteins and lipids. In addition, the DNA strands renature at neutral pH. The chromosomal DNA, which is much longer than the plasmid DNA, renatures as a tangle that gets trapped in the SDS precipitate. The plasmid DNA renatures normally and stays in solution, effectively separating plasmid DNA from the chromosomal DNA and the proteins and lipids of the cell.
Normally in 20.109 we do an in-house mini-prep procedure according to the steps above followed by ethanol precipitation. However, because you are isolating so many colonies, today we will use a commercially available kit so that the work can go more quickly. The principle is the same as that of our "quick and dirty" (and cheaper!) prep, but is combined with the silica gel column purification you are familiar with from using other Qiagen kits.
After isolation, you will quantify your DNA by spectrophotometry. Nucleic acids (both RNA and DNA) have an absorbance peak at 260 nm. Beer's law may be used to quantify the amount of DNA from this peak: Abs = ε l c, where Abs is the measured absorbance, l is the path length (1 cm for most specs), c is concentration, and ε is the extinction coefficient. For DNA, ε is 0.02 (μg/mL cm)-1, so 1 absorbance unit corresponds to 50 μg/mL of DNA. The absorbance at 280 nm gives some indication of DNA purity, as proteins have their absorbance peaks at that value (primarily due to the aromatic peptides tryptophan and tyrosine). An Abs260:Abs280 ratio of ~1.8:1 is desired.
Miniprepped DNA will be sent for sequencing off-site; each clone will be sequenced from two directions, in order to cover the entire 1400 bp region of interest. Together, you'll do 256 sequencing reactions per lab section!
Normal bases versus chain-terminating bases
The invention of automated sequencing machines has made sequence determination a relatively fast and inexpensive endeavor. The method for sequencing DNA is not new but automation of the process is recent, developed in conjunction with the massive genome sequencing efforts of the 1990s. At the heart of sequencing reactions is chemistry worked out by Fred Sanger in the 1970s which uses dideoxynucleotides (see schematic above left). These chain-terminating bases can be added to a growing chain of DNA but cannot be further extended. Performing four reactions, each with a different chain-terminating base, generates fragments of different lengths ending at G, A, T, or C. The fragments, once separated by size, reflect the DNA’s sequence. In the “old days” (all of 15-20 years ago!) radioactive material was incorporated into the elongating DNA fragments so they could be visualized on X-ray film (image above center). More recently fluorescent dyes, one color linked to each dideoxy-base, have been used instead. The four colored fragments can be passed through capillaries to a computer that can read the output and trace the color intensities detected (image above right). Your sample was sequenced in this way on an ABI 3730 DNA Analyzer.
Analysis of sequence data is no small task. “Sequence gazing” can swallow hours of time with little or no results. There are also many web-based programs to decipher patterns. The nucleotide or its translated protein can be examined in this way. Thanks to the genome sequence information that is now available, a new verb, “to BLAST,” has been coined to describe the comparison of your own sequence to sequences from other organisms. BLAST is an acronym for Basic Local Alignment Search Tool, and can be accessed through the National Center for Biotechnology Information (NCBI) home page.
In another week you will finally get to see the results of all your hard work...
Part 1: Extract DNA from selected clones (mini prep)
Each partner today has eight minipreps to do. You may find it easier to complete the sixteen total minipreps in two shifts than to attempt to "quickly" pipet across sixteen samples. One approach you could take is the following: start with the first eight candidates, then spin down the next eight during the 5 min lysis step, and finally start adding P1 to the second batch of eight when you reach the 10 min spin step for the first batch.
- Pick up your eight candidates cultures, which are growing in the test tubes labeled with your team color. Label eight eppendorf tubes to reflect your candidates (C1-8).
- Vortex the bacteria and pour ~1.5 mL of each candidate into an eppendorf tube.
- Balance the tubes in the microfuge, spin them at maximum speed for two minutes, and remove the supernatants with the vacuum aspirator.
- Pour another 1.5 mL of culture onto the pellet, and repeat the spin step.
- Resuspend the cell pellet in 250 μL buffer P1.
- Buffer P1 contains RNase so that we collect only our nucleic acid of interest, DNA.
- Add 250 μL of buffer P2 and mix by inversion until the suspension is a homogeneous blue color. About 4-6 inversions of the tube should suffice. You may incubate here for up to 5 minutes, but not more.
- Buffer P2 contains sodium hydroxide for lysing.
- The blue color comes from a special reagent that is not required for purification, but is simply used to check one's mixing technique.
- Add 350 μL buffer N3, and mix immediately by inversion until there is no blue colour (4-10 times).
- Buffer N3 contains acetic acid, which will cause the chromosomal DNA to messily precipitate; the faster you invert, the more homogeneous the precipitation will be.
- Buffer N3 also contains a chaotropic salt in preparation for the silica column purification.
- Centrifuge for 10 minutes at maximum speed. Note that you will be saving the supernatant after this step.
- Meanwhile, prepare 8 labeled QIAprep columns, one for each candidate clone, and 8 trimmed eppendorf tubes for the final elution step.
- Transfer the entire supernatant to the column and centrifuge for 1 min.
- Wash with 0.5 mL PB, then separately with 0.75 mL PE, with each spin step 1 min long.
- After removing the PE, spin the mostly dry column for 1 more min.
- It is important to remove all traces of ethanol, as they may interfere with subsequent work with the DNA.
- Add 30 μL of buffer EB to the top center of the column, wait 1 min, and then spin 1 min to collect your DNA.
Part 2: Measure DNA concentration
- For each DNA sample, you will add X μL of miniprepped DNA to Y μL of water. This ought to give you a measurement that's in a reliable range. You can prepare labeled eppendorf tubes containing the appropriate amount of water in advance.
- To get the most accurate measurement, you should also prepare a blanking solution that contains everything except the RNA, i.e., Y μL of water and X μL of EB elution buffer.
- Add Y- μL of your eight solutions (per person) to separate cuvettes and head to the spec.
- Notice that today we are using cuvettes made of a special plastic that is transparent to UV light.
- Enter DNA measurement mode [EXACT NAME??]
- Begin with the cuvette containing blanking solution, and hit Blank on the spectrophotometer.
- Proceed to take an absorbance scan of each DNA sample. Record the 260 nm and 280 nm absorbance values in your notebook. The spec will calculate the purity ratio for you, but not the DNA concentration.
- What assumption are we making about the cuvettes when we put the blanking solution in a separate cuvette from the samples?
- Calculate the eight DNA concentrations, along with the volume of DNA required to use 500 ng in a sequencing reaction. You may find the table below helpful.
- Recall that an absorbance value of 1 indicates a concentration of 50 μg/mL of the measured DNA (i.e., the diluted solution and not the original stock).
- Which of the two wavelengths indicates DNA quantity? Re-read the introduction if you're not sure.
- If any of your samples requires adding less than 1-2 μL of DNA, prepare an intermediate dilution first. For example, instead of adding 0.9 μL of stock, you could add 9 μL of a 1:10 dilution of that particular miniprep. No sequencing reaction should contain more than 10 μL of DNA.
Part 3: Prepare sequencing reactions
As we will discuss in lab today, sequencing reactions require a primer for initiation. Legible readout of the gene typically begins about 40-50 bp downstream of the primer site, and continues for ~1000 bp at most. Thus, multiple primers must be used to fully view genes > 1 Kbp in size, such as your ~ 1400 bp 16S sequence. We will use forward and reverse primers that anneal to the vector upstream and downstream of the 16S insert in order to capture the entire sequence for analysis.
The recommended composition of sequencing reactions is ~500 ng (CHECK) of plasmid DNA and 25 pmoles of sequencing primer in a final volume of 15 μL. Each of your sequencing reactions should thus consist of 5 μL of primer stock, and 10 μL of aqueous DNA. (For example, if you are adding only 2 μL of DNA for a particular candidate, you must add another 8 μL of water to complete the sequencing mixture.)
Each person will fill in 2 columns on a 96-well plate, according to the table below...
Part 4: Begin sensitivity/specificity analysis for microsporidia primers
For each team, V corneae and/or E hellem templates were amplified according to the tables below. On a gel, ...
Part 5: Count colonies
When you have a moment today, separately count the white, the light blue, and the blue colonies on your plate from last time and post them on today's Talk page. Note that light blue colonies may contain insert, particularly if the insert leaves the entire complementation peptide in-frame.
Don't forget to count the colonies we have already picked! These colonies are indicated by the starting numbers in the table.
For next time
Some of you have journal clubs next time. No other homework is due on Day 6. There is homework due for everyone on Day 7.
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