Characterization of Microbial Communities (or Isolates) using 16S Amplification
Isolate Genomic DNA From Soil Sample
Protocol for Using the Power® Soil DNA Isolation Kit by Mo Bio Laboratories'
Manufacturer Information at 
Please wear gloves during this protocol
1. You will each use one sample that you froze after collection at WE Farm. Transfer ALL of the sample from the microfuge tube to a PowerBead Tube and label this tube on the top only with your initials and add a piece of your team color tape around the tube.
2. Gently vortex to mix.
3. Check Solution C1 to see that it's not precipitated. If Solution C1 is precipitated, heat solution to 60C until the precipitate has dissolved before use.
4. Add 60 microliters of Solution C1 to your PowerBead tube and invert several times or vortex briefly.
5. Give your labeled PowerBead tube to your instructor who will take all the PowerBead Tubes to a power vortex in the 4°C coldroom. Your samples will "turbovortex" for 7 min at full speed. Alternatively, we can use a FastPrep® Bead Beater for 45 seconds at speed 5 (or full speed. Note: only 2-3 samples will fit in the Fast Prep® BeadBeater at a time.
What’s happening: The BeadBeating step is critical for complete homogenization and cell lysis. Cells are lysed by a combination of chemical agents from steps 1-4 and mechanical shaking introduced at this step. By randomly shaking the beads in the presence of disruption agents, collision of the beads with microbial cells will cause the cells to break open.
6. Microcentrifuge your tubes at 10,000rcf for 1 minute at room temperature. Caution: Be sure not to exceed 10,000rcf or the tubes may break. Make sure the PowerBead tubes rotate freely in your centrifuge without rubbing!
7. Transfer the supernatant (don't transfer the beads!) to a clean 2 ml Collection Tube. If you don't know what a collection tube is, ask your instructor. Don't use a regular microfuge tube.
Note: Expect between 400 to 500 microliters of supernatant at this step. The exact recovered volume depends on the absorbancy of your starting material and is not critical for the procedure to be effective. The supernatant may be dark in appearance and still contain some particles. The presence of carry over or a dark color in the mixture is expected in many sample types at this step.
Subsequent steps in the protocol will remove both carry over soil and coloration of the mixture.
8. Add 250 microliters of Solution C2 to the collection tube and vortex for 5 seconds. Incubate at 4C for 5 minutes.
What’s happening: Solution C2 contains a patented reagent to precipitate non-DNA organic and inorganic material including humic substances, cell debris, and proteins. It is important to remove contaminating organic and inorganic matter that may reduce DNA purity and inhibit downstream DNA applications.
9. Centrifuge the Collection Tube at room temperature for 1 minute at 10,000rcf.
10. Avoiding the pellet, transfer up to, but no more, than 600 microliters of supernatant to a clean 2 ml Collection Tube (provided).
What’s happening: The pellet at this point contains non-DNA organic and inorganic material including humic acid, cell debris, and proteins. For the best DNA yields, and quality, avoid transferring any of the pellet.
11. Add 200 microliters of Solution C3 and vortex briefly. Incubate at 4C for 5 minutes.
What’s happening: Solution C3 is a second reagent (patented) to precipitate additional non-DNA organic and inorganic material including humic acid, cell debris, and proteins. It is important to remove contaminating organic and inorganic matter that may reduce DNA purity and inhibit downstream DNA applications.
12. Centrifuge the tube at room temperature for 1 minute at 10,000rcf.
13. Avoiding the pellet, transfer up to, but no more, than 750 microliters of supernatant to a clean 2 ml Collection Tube (provided).
What’s happening: The pellet at this point contains additional non-DNA organic and inorganic material including humic acid, cell debris, and proteins. For the best DNA yields, and quality, avoid transferring any of the pellet.
14. Shake to mix Solution C4 before use. Add 1.2 ml (do this by adding 600 microliters twice) of Solution C4 to the supernatant (be careful solution doesn’t exceed rim of tube) and vortex for 5 seconds.
What’s happening: Solution C4 has a high concentration of salts. Since DNA binds tightly to silica at high salt concentrations, this will adjust the DNA solution salt concentrations to allow binding of DNA, but not non-DNA organic and inorganic material that may still be present at low levels, to the Spin Filters.
15. Load approximately 675 microliters of the C4 + supernatant mixture from the previous step onto a Spin Filter sitting in a Collection Tube (save the remainder of the supernatant!!) and centrifuge the spin filter at 10,000rcf for 1 minute at room temperature. Discard the flow through (NOT the spin filter!!!) and put the spin filter back in the Collection Tube. Add an additional 675 microliters of the Step 14 mixture to the same Spin Filter and centrifuge at 10,000rcf for 1 minute at room temperature. Discard the flow through and load the remainder of the Step 14 mixture onto the Spin Filter in the Collection Tube and centrifuge at 10,000rcf for 1 minute at room temperature.
Note: A total of three loads for each sample processed are required. You will using the same Spin Filter and Collection Tube for all 3 spins.
What’s happening: DNA is selectively bound to the silica membrane in the Spin Filter device in the high salt solution. Contaminants pass through the filter membrane, leaving only DNA bound to the membrane.
16. Add 500 microliters of Solution C5 to the Spin Filter in the Collection Tube and centrifuge at room temperature for 30 seconds at 10,000rcf.
What’s happening: Solution C5 is an ethanol based wash solution used to further clean the DNA that is bound to the silica filter membrane in the Spin Filter. This wash solution removes residual salt, humic acid, and other contaminants while allowing the DNA to stay bound to the silica membrane.
17. Discard the flow through (not the Spin Filter) from the 2 ml Collection Tube.
What’s happening: This flow through fraction is just non-DNA organic and inorganic waste removed from the silica Spin Filter membrane by the ethanol wash solution.
18. Centrifuge the Spin Filter in the Collection Tube again at room temperature for 1 minute at 10,000rcf.
What’s happening: This second spin removes residual Solution C5 (ethanol wash solution). It is critical to remove all traces of wash solution because the ethanol in Solution C5 can interfere with many downstream DNA applications such as PCR, restriction digests, and gel electrophoresis.
19. Carefully place Spin Filter in a clean 2 ml Collection Tube (provided). DO NOT transfer any liquid that may be on the bottom of the spin filter basket and avoid splashing any Solution C5 onto the Spin Filter.
Note: It is important to avoid any traces of the ethanol based wash solution in the elution that will be created in the next step.
20. Add 100 microliters of Solution C6 to the center of the white Spin Filter membrane.
Note: Placing the Solution C6 (sterile elution buffer) in the center of the small white membrane will make sure the entire membrane is wetted. This will result in a more efficient and complete release of the DNA from the silica Spin Filter membrane. As Solution C6 (elution buffer) passes through the silica membrane, DNA that was bound in the presence of high salt is selectively released by Solution C6 (10 mM Tris) which lacks salt.
Alternatively, sterile DNA-Free PCR Grade Water may be used for DNA elution from the silica Spin Filter membrane at this step (MO BIO Catalog# 17000-10). Solution C6 contains no EDTA. If DNA degradation is a concern, Sterile TE may also be used instead of Solution C6 for elution of DNA from the Spin Filter.
21. Centrifuge the Spin Filter in its Collection Tube at room temperature for 30 seconds at 10,000rcf.
22. Discard the Spin Filter. The DNA in the collection tube is now eluted and ready for freezing or for use as a PCR template.
DNA is eluted in Solution C6 (10 mM Tris) and must be used immediately or stored at -20to -80C to prevent degradation.
Make sure your DNA is labeled. We need to measure the DNA concentration before we freeze your extract. DNA at 100ng/microliter is desirable for the next step (amplification of 16s rDNA by polymerase chain reaction) in our culture independent approach to understanding the make up of your soil community. Once you know the concentrate of your DNA, you can adjust the volume of DNA template used in the pcr reaction. The directions for determining the DNA concentration in your extract is described below.
The genomic DNA extraction has, no doubt, resulted in a mixed DNA population from a myriad of microorganisms and DNA from plants, insects, or other multi-cellular life forms in the community. Since we are most interested in the scope of our bacterial population, we will amplify, by polymerase chain reaction, only bacterial DNA 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 20 cycles of polymerase chain reaction in a thermal cycler, the result will be a pcr product containing hundreds, if not thousands, of unique copies of 16s rDNA. This amplification of exclusively bacterial DNA will allow identification of much of the bacterial flora present in the community, most of which is unculturable by conventional techniques.
Part A: 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
Obain 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 sample. Do not use tape.
If your DNA is at approximately 100ng/μL, you will follow the Template Table (shown below) adding 3μL DNAase free water and only 1μL of template DNA to the reagents that have already been premixed for you in your pcr tube (10μL master mix, 4μL DNAase free water, 1μL of each of 2 primers).
If your DNA concentration was less than 20ng/μL, you will add 4 μL of DNA and no extra water. If your concentration was between 20 and 100ng/μL, calculate how much template DNA to add by using the formula 100 / your isolate's DNA conc. Add that number of microliters of DNA (not more than 4) and enough DNAase free water so that the number of microliters of DNA + microliters of water =4. Example: Your DNA conc. was 33ng/μL. 100/33 = 3.3 so you would add 3.3μL of DNA and 0.7μL of DNAase free water. 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 other liquid ingredents, 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.
| Component || amt. in a 20 μl|
| Final Conc.
| 4 μL already in tube.|
Want to achieve
total of 20 μl reaction vol.
Add from 0 - 3μl
| 2x Phusion Master Mix
|| 10 μl
| 27F primer
|| 0.5 μMolar
| 1492R primer
|| 0.5 μMolar
| template DNA
|| 1-4 μl
|| optimum is 100ng of DNA/reaction
The cycling program is shown below.
Thermal Cycler Program:
3 step program
| Cycle Step || Temperature || Time || # of Cycles
| Initial Denaturation
|| 5 min.
| Denaturation |
| 98C |
| 10 sec |
| Final Extension
|| 72C |
| 10 min |
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 cloning next week are found later in this lab description. 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: Clean Up of pcr product using Epoch BIoLabs GenCatch PCR CleanUp Kit
Before we can ligate our bacterial 16s rDNA 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
Part C: 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 to the data folder in the First Class lab conference 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 100bp 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!
Make sure your pcr product is clearly labeled as pcr product and has your initials, team color, lab section (Tues or Wed), soil identifier code. Your instructor will measure the new DNA conc. using the nanodropper 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 rDNA genes from your soil bacteria community into a special genetically engineered cloning vector. Once the a soil 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 rDNA 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.