Creating Lysates from your Isolates
One of the big advantages of the molecular revolution in microbial ecology is that we can actually find, based on 16S rRNA gene sequences, the identity of your isolates. In order to do that, we will follow the protocol below which first involves liberating the DNA in your isolate, then amplifying the 16S rRNA gene, then sequencing it. The bioinformatics analysis of this sequence will be covered next time we meet.
Isolate Genomic DNA From Your Sample
Please wear gloves during this protocol
1. For each of your isolates, label a 200 ul PCR tube and fill it with 100 ul of water.
2. Using a sterile loop, transfer a small bit (barely visible) of your isolate into the 100 ul solution in the PCR tube - make sure to shake the isolate off the loop so that it is in the water
3. Cap and vortex the PCR tube for 10 seconds.
4. Place in the PCR machine at 95C for 10 minutes.
Because you've just made a lysate of your isolate, remember that this solution contains all cellular contents, including DNAses -- enzymes that break down that very DNA we are trying to amplify. You can store your lysates at -20C in the freezer in the lab but make sure to create new ones if you freeze-thaw them more than once.
We will next attempt to amplify only bacterial rRNA genes by using "universal" bacterial primers :a forward primer, Eub27F (5′–3′:AGA GTT TGA TCC TGG CTC AG) , and a reverse primer, Eub1492R (5′–3′: ACG GCT ACC TTG TTA CGA CTT). These primers are short sequences of single stranded DNA that are complementary in sequence to areas of the 16s rRNA gene. The 16S rRNA gene sequence is particularly good target gene for amplification because this gene (encoding a ribosomal subunit) contains conserved sequences of DNA common to all bacteria (to which the primers are directed) as well as divergent sequences unique to each species of bacteria (allowing identification of the bacterial species from sequence databases and sequence identifying software). Our "universal" primers will anneal to most bacterial DNA and initiate an exponential amplification of the 16s rRNA gene from the template DNA. After 30 cycles of polymerase chain reaction in a thermal cycler, the result will be a pcr product containing hundreds, if not thousands, of the 16s rRNA gene.
PCR Amplification of 16s rRNA genes from Universal Bacterial Primers
To review how the polymerase chain reaction works and how it exponentially amplifies specific sequences of DNA, go to the following web site:
All PCR reactions require a thermal cycler to elevate and reduce the reaction temperature quickly and keep it at a specific temperature for a prescribed amount of time. There is a basic pattern to these temp. cycles, but there are differences, so you must be sure to program the cycler with the correct time and temperature for your specific amplification. Traditionally, pcr used Taq polymerase, a heat stable DNA polymerase originally found in a extremophilic bacterium, Thermus aquaticus, that lives and reproduces in boiling hot springs. We are not using Taq for our pcr but a different polymerase, Finnzyme's Phusion High-Fidelity Polymerase, a proprietary reagent that uses a novel heat-stable Pyrococcus-like enzyme. Phusion DNA Polymerase generates long templates with a greater accuracy and speed than with Taq. The error rate of Phusion DNA Polymerase in Phusion HF Buffer is determined to be 4.4 x 10-7, which is approximately 50-fold lower than that of Thermus aquaticus DNA polymerase, and 6-fold lower than that of Pyrococcus furiosus, another proof-reading DNA polymerase.
Therefore, our pcr product DNA will have far fewer "mistakes" in the sequences that are replicated from template DNA. Our polymerase will also work much faster so our ~20 cycles will require less time than conventional Taq based pcr.
Protocol for PCR
Obtain a tiny 0.2ml pcr tube from your instructor. All of the ingredients listed below in the table, except the template DNA, have been added together previously and kept on ice for you in these tubes.
Label it with a fine tipped Sharpie on the top and side with the code name for your isolate. Do not use tape.
you will add 4 μL of lysate to your PCR tube with master mix. Since your pcr tube already has 10μL master mix, 4μL DNAase free water, and 1μL of each of 2 primers, the total reaction volume for everyone will be 20μL.
It is very important to pipet these tiny volumes accurately. Use the P10 or P20 pipettes. Look at the tip after you draw up your measured volume to make sure you have liquid there.
Dispense the template DNA into the liquid directly, watching to make sure that the liquid has left the pipette tip.
Tap the bottom of the tube (VERY GENTLY!) and flick the tube to mix. Do not treat these tubes roughly as they are quite thin-walled and can break or crack.
Bring your tube to your instructor; they will show you where the thermal cycler is located in JH 022. Your instructor will start the reaction when everyone's tubes are loaded.
||amt. in a 20 μl
|4 μL already in tube.
Want to achieve
total of 20 μl reaction vol.
Add from 0 - 3μl
|2x Phusion Master Mix
||optimum is 100ng of DNA/reaction
The cycling program is shown below.
Thermal Cycler Program:
3 step program
||# of Cycles
While the 16S rRNA genes from all of the bacterial species in your DNA are being amplified in the thermal cycler, you will have about an hour to work on any other parts of your project.
After the PCR reactions are complete, you will need to complete a "Clean-Up" of your pcr products (remove the unused dNPTs, primer dimers, salts, etc. The instructions for using a kit to purify your pcr products and get them ready for sequencing next time. You will also need to set up a gel to assess the purity of your pcr product and the success of your amplification.
Part B: Agarose Gel Electrophoresis of Clean PCR PRODUCT
To see if you successfully amplified the 16s rRNA gene and not anything else, you will "run a gel" on your cleaned pcr products. To run a gel means that we will perform an electrophoretic separation of the DNA fragments in your cleaned up pcr product, using 1/10 vol. of your pcr product applied to a 1% agarose gel stained with Sybr Safe DNA stain. Your instructor will photograph the gel, label it with your amplicon id from the template and post the gel photo on OnCourse so you can evaluate your success at 16S rRNA gene amplification. You should see a single band of ~1.5kb indicating that the only dsDNA in your pcr product came from amplification of a ~1500bp gene fragment. Can you explain how we know the size of our amplified gene fragment?
Your agarose gel is made of 1.0% agarose (w/v) in 1x TBE buffer (10x=890mM Tris, 890mM Boric Acid, 20mM EDTA) with SybrSafe™ stain.
DNA is uniformly negatively charged and will,therefore, move toward the positive electrode. The separation is determined by the size or mass of the molecule or fragments of DNA.
Procedure for Agarose Gel Electrophoresis of PCR products
Load 1/10 of the total volume of pcr product (1 microliter minimum). In our case we should load 5 microliters.
You will put the 5 microliters of your pcr product as a spot on a small piece of parafilm and add 5 microliters of loading dye (0.25% XC, 30% glycerol, 0.1mg/ml RNAase). Mix the loading dye by pipetting up and down before loading all 10 microliters into a lane of the 1% agarose gel (1% wt/vol in 1xTBE buffer with Sybr Safe DNA stain (a proprietary reagent from Invitrogen used according to manufacturer's directions at http://www.invitrogen.com). Record on the gel template in which well you have loaded your pcr product. Be sure to leave the first two lanes and the last lane empty for the 1kbp ladder, the positive control and the negative water control.
Note that Loading dye contains glycerol to keep our sample in the lane rather than floating away and will have one of 3 marker dyes (bromophenol blue, xylene cyanol, or orange G) that facilitate estimation of DNA migration distance and optimization of agarose gel run time. 1x TBE buffer is used in this electrophoretic separation (89mM Tris, 89mM Boric acid, 2.0mM EDTA. The gel will be run at 120V for approximately 30 minutes.
How will you judge a successful amplification? How many fragments and of what size do you expect to see?
Make sure you give back the rest of your soil DNA isolate and the rest of the cleaned up pcr product to your instructor to freeze after the gel is loaded. Both are now in identical looking microfuge tubes with volume being the only visible difference. Make sure it is clear which is the pcr product and which is the genomic DNA isolate!
Part C: Clean Up of pcr product using Epoch BIoLabs GenCatch PCR CleanUp Kit
Before we can ligate our bacterial 16s rRNA genes into vector plasmids, we must remove interfering dNPTs, primers, and other small degraded DNA. We will use a column that separates DNA by size. Since the reagents and column materials in the kit we will use are proprietary, we won't know exactly what is going on at each step but, basicially, we will apply our pcr product to a column of a particular density, wash away elements too small to be trapped in it, and elute off the larger fragments of DNA (that should be ~1500bps if our pcr amplification of the 16s rRNA genes in our soil genomic DNA was successful).
Notes before Starting:
95% ethanol has been added to Buffer WS before first time use (see bottle label for volume).
All centrifuge steps are carried out at 17,900rfc (~13,000 rpm in a microcentrifuge) in a conventional tabletop microcentrifuge at room temperature.
1. Measure 500 μl of Buffer PX using your P1000 and add part of it to your pcr product and the rest to a clean microfuge tube. Using your P200 set to 200 μL, remove all the pcrProduct/buffer mix in the pcr tube and add it to the PX buffer in the microfuge tube. Close the cap of the microfuge tube and mix.
2. Place a GenCatch™ spin column in a provided 2 ml collection tube.
3. Load all of the pcr product/bufferPX mixture created in step 1 (up to a maximum of 700μL total volume) to the spin column and centrifuge for 60 sec.
4. Discard flow-through. Place the spin column back into the same collection tube.
(Collection tubes are re-used to reduce plastic waste.)
5. If you applied all the pcr product to the spin column in step 3, skip this step and proceed to step 6. If you had more than 700 μL volume of pcrProduct/bufferPX made in step 1, apply the remaining volume to the spin column and centrifuge for 1 minute. Discard the flow-through and place the spin column back in the same collection tube.
6. Wash the spin column by adding 500 μL Buffer WF to the spin column and centrifuge for 60 sec. Be careful to use WF buffer!!
7. Discard flow-through and place the spin column back in the same collection tube.
8. Wash the spin column by applying 700 μL of Buffer WS. Note that WS Buffer is different than the buffer used in step 6!!! Centrifuge the column for an additional 1 min. Check that ALL the buffer is in the flow-through, if there is buffer remaining in the spin-column, re-spin if needed. Discard the flow-through.
9. Centrifuge the spin-column in the same collection tube at full speed for 3 more minutes to remove ALL ethanol residue. It is crucially important to remove all ethanol residue; residual ethanol may inhibit subsequent enzymatic reactions.
10. Place each spin column into a new, clean 1.5 ml microcentrifuge tube (not a collection tube).
11. To elute DNA, add 50μl of the Elution Buffer EB (10 mM Tris·Cl, pH 8.5) to the center of each spin column membrane. Let it stand for 2 minutes to allow it completely adsorb and then centrifuge the spin column in the microfuge tube for 1 min at 17,900 x g (13,000 rpm).
Keep your pcr product on ice until your instructor tells you that it's time to load the gel in order to determine the success of this amplification and clean-up.
IMPORTANT NOTES for using this kit: Ensure that the elution buffer (EB) is dispensed directly onto the spin column membrane for complete elution of bound DNA. The average eluate volume is 48 μl from 50 μl elution buffer volume.
Elution efficiency is dependent on pH. The maximum elution efficiency is achieved
between pH 7.0 and 8.5. Store DNA at –20°C as DNA may degrade in the absence of a buffering
Make sure your pcr product is clearly labeled. Your instructor will measure the new DNA conc. using the nanodrop and post those concentrations for you in ng/μL. In the next lab you will use the most successful of your team's pcr amplifications of 16s rDNA and use those pcr products to ligate the 16s rRNA gene genes from your bacteria community into a special genetically engineered cloning vector. Once the a bacterium's 16s rRNA gene is incorporated into those vector plasmids, we will transform competent genetically engineered E. coli bacteria with a plasmid. Transformants (bacteria that have taken up a vector plasmid and express its genes) will be plated on selective media to find cells containing the 16s rRNA gene insert. Eventually we will send away some of those E. coli for sequence analysis to determine the identity of some of the bacterial community members in your original soil sample.
Part 1: Culture-Independent Identification of Soil Bacteria
Your instructor will return your frozen, cleaned-up pcr products containing amplified fragments of 16s rRNA gene from many of the species of soil bacteria in your soil sample. Today you will insert your bacterial 16s rRNA gene fragments into a patented cloning vector (pCR-BluntII TOPO®) and then transform that vector into a special genetically engineered strain of Escherichia coli bacteria that will express a vector gene for kanamycin resistance, allowing us to select for transformants on media containing kanamycin.
The principle behind TOPO® cloning is the enzyme DNA topoisomerase I, which will function in this system both as a restriction enzyme and as a ligase. Its biological role is to cleave and rejoin DNA during replication. Vaccinia virus topoisomerase I specifically recognizes the pentameric sequence 5´-(C/T)CCTT-3´ and forms a covalent bond with the
phosphate group attached to the 3´ thymidine. It cleaves one DNA strand, enabling the DNA to unwind. The enzyme then religates the ends of the cleaved strand and releases itself from the DNA. To harness the religating activity of topoisomerase, TOPO® vectors are provided linearized with topoisomerase I covalently bound to each 3´ phosphate. This enables the vectors to quickly ligate DNA sequences with compatible ends.
We used a polymerase that creates blunt ended DNA fragments rather than using TaQ. Taq polymerase makes fragments with 3' T overhangs; therefore, complementary single stranded A rich "sticky ends" allow ligation. Blunt ends require a different Blunt-fragment cloning protocol. Invitrogen's Zero Blunt® TOPO® PCR Cloning Kit will work well for us. It has several (T7, SP6, and M13 forward and reverse) priming sites for directing sequencing to the appropriate region and it has two resistance genes, Kanamycin and Zeocin, for selecting clones in a genetically engineered form of E. coli that we will use for separating the amplified 16s rRNA genes from our soil flora.
Additionally, the cloning system we will use contains two different background reducers, one of which is a lethal ccdB (control of cell death)gene encoding a ccdB protein that poisons bacterial DNA gyrase, causing degradation of the host chromosome and cell death. When one of our 16s rRNA genes from our pcr product is ligated into the vector, the ccdB gene is disrupted, enabling recombinant colonies to grow while other colonies without a vector insert will not grow. Because a few colonies may form despite the undisrupted expression of ccdB there is a second mechanism of insuring that we only pick colonies coming from cells with our 16s rRNA gene insert. As added insurance that we will select only colonies that are transformed with a plasmid vector with a 16s rRNA gene insert, there is a lacZ gene positioned next to the ccdB gene in the vector. LacZ encodes beta-galactosidase, an enzyme that catalyzes the breakdown of colorless substrates such as Xgal (5-Bromo-4-chloro-3-indolyl beta-Dgalactopyranoside) to a colored cleavage product (in this case, a blue product). However, the promoter for transcription of the ccdB gene AND the lacZ gene is disrupted by the insertion of the 16s DNA insert. Because of this disruption of transcription regulation, the lacZ gene product (beta-galactosidase) and the ccdB product (gyrase poison)are not produced in appreciable quantity. Colonies that are transformed with "empty" vectors will be differentiated visually by color from those that contain our 16s rRNA gene insert on media with X-gal. Cells containing a plasmid vector with our 16s RNA gene have disruption of both LacZ and ccdB gene regulation. They will not be killed by absence of DNA gyrase and those colonies will be white. They will live and form "not-blue" colonies because the Xgal in the medium will not be converted to a blue product due to lack of the catalzying enzyme, beta-galactosidase. You will look for white or "not-blue" colonies. (Cool technology!)
Part A: Using Zero Blunt TOPO PCR Cloning Kit with One Shot TOP 10 Chemically Competent E. coli
PCR cloning requires three steps.
We will clone three pcr products/per sampling site, if your team had three successful amplifications. If you had 4 successful amplifications from your sampling site, use the most successful 16s rRNA gene amplifications and omit the weakest one.
Procedure: Add the reagents in this order!
1. Add 2 μl of PCR product to a 0.2ml pcr tube (your team color)
2. Add 1 μL of salt solution (final conc. 200mM NaCl, 10mM MgCl2).
3. Add 2 μL of purified HPLC water (DNAase free).
4. Add 1 μL of pCR®II-Blunt-TOPO® cloning vector plasmid. (MAKE sure you pipet this correctly with a P2 and a filter tip!)
4. Incubate 15 min at room temperature.
5. Continue to next step: Transform Oneshot Top10 competent E. coli.
Part B Transforming TOPO Competent E. coli
Genotype of OneShot TOP10 Competent Cells: F- mcrA Δ(mrr-hsdRMS-mcrBC) φ80lacZΔM15 ΔlacX74 recA1 araD139 Δ(araleu) 7697 galU galK rpsL (StrR) endA1 nupG
General Handling: Be extremely gentle when working with competent cells. Competent cells have been chemically treated to make their cell walls and membranes more porous so they are fragile and highly sensitive to changes in temperature. They can be easily lysed by too vigorous pipetting. Transformation should be started immediately following the thawing of the cells on ice. Mix by swirling or tapping the tube gently, not by pipetting(no vortexing).
Transforming One Shot® Competent Cells
Introduction: Once you have performed the TOPO® Cloning reaction, you will transform your pCR®-Blunt II-TOPO® construct into TOPO10 competent E. coli provided with your kit.
You will need the following reagents and equipment:
• TOPO® Cloning reaction from Performing the TOPO® Cloning Reaction
• S.O.C. medium (super optimal broth medium:0.5% Yeast Extract;2% Tryptone;10 mM NaCl;2.5 mM KCl;10 mM MgCl2;10 mM MgSO4;20 mM Glucose)This medium is included with the kit)
• 42°C water bath or heat block
• WARM Luria-Bertoni (LB) solid medium containing 50 μg/ml kanamycin and 50μL/ml Xgal (5-Bromo-4-chloro-3-indolyl beta-Dgalactopyranoside)
• 37°C shaking and non-shaking incubators
Preparing for Transformation
For each transformation, you will need one vial of competent cells and two
selective medium agar plates.
• Equilibrate a water bath to 42°C
• Bring the vial of S.O.C. medium to room temperature.
• Warm LB plates containing 50 μg/ml kanamycin and Xgal at 37°C
for 30 minutes.
• Thaw on ice 1 vial of One Shot® cells for each transformation.Don't remove them from the -80C until ready for use.
1. Add 2 μl of the TOPO® Cloning reaction when it is completed into a vial of One Shot® Chemically Competent E. coli and mix gently by swirling. Do not mix by pipetting up and down!
2. Incubate on ice for 10 minutes.
Note: Longer incubations on ice do not seem to have any affect on transformation
efficiency. The length of the incubation is at the user’s discretion.
3. Heat-shock the cells for 30 seconds exactly at 42°C in the heatblock (without shaking).
4. Immediately (take your ice bucket with you to the heat block) transfer the tubes to ice .
5. Add 250 μl of room temperature S.O.C. medium (it must NOT be cold).
6. Cap the tube tightly and put the capped tube in a empty non-sterile 15 ml. conical tube and shake the tube horizontally (200 rpm) at 37°C for
1 hour. While the shaking is going on, slightly dehydrate 2 LB + kan + Xgal plates by placing them with lids slightly agar in the laminar flow hood with the blower on for 10 min. Then place the plates in the 37C incubator to prewarm. (The plates must NOT be cold when transformed cells are plated.)
7. After the 1 hour incubation of the transformation mix, Use your P200 micropipet to pipet 50 μl from each transformation to the center of a prewarmed LB + kan+ Xgal plate. Using a disposable sterile plastic spreader, carefully spread the aliquot of cells over the entire surface of the plate.
8. Repeat step 7 on a new LB + kan + Xgal plate, using a 200 μL volume of transformed cells. You will plate two different volumes to ensure that at least one plate will have well-spaced colonies.
9. Incubate all plates upside down overnight at 37°C. Remember to label each plate with all the appropriate information: your initials, lab section, date, your soil sample id, the type of medium, and the id of the cells and volume used. Refrigerate the remainder of your transformed cells at 4C overnight in case you need to plate a smaller number of cells to achieve isolated colonies. Check your transformations after 12-18 hours (overnight incubation)to be sure of successful transformation. When medium size, ISOLATED colonies, have appeared, refrigerate your transformation plates until LAB 5. DO NOT LEAVE THEM INCUBATING TOO LONG, resulting in overgrown colonies that are not isolated! If you have no transformation or a lawn of growth after the initial overnight incubation, contact your instructor immediately for help. You will need to reisolate by plating a diluted or smaller volume of cells on a new plate or redo the cloning and transformation if none of the transformations from your soil community is successful.
10. An efficient TOPO® Cloning reaction will produce several hundred
colonies. The colonies with inserts will be white or, at least, "not-blue". Look at the map of the cloning vector and the background information description of the cloning and figure out why all colonies should have soil genomic 16s rRNA inserts and why those that are not blue are particularly likely to be the ones we want.