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:
PCR animation http://www.dnalc.org/resources/animations/pcr.html
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 (choose the one prepared for your team in your team's color). 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 soil DNA isolate 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 soil isolate 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 a P2 or P10 and the special small tips with a filter when pipetting DNA. Look at the tip after you draw up your measured volume to make sure you have liquid there.
Dispense the template DNA onto the side wall of the pcr tube close to the other liquid ingredents, watching to make sure that a small bead of liquid is left on the wall of the tube.
Without changing the tip, pipet up and down in the pcr mix to wash the tip and then wash some of the mixture over the bead of template DNA that may still be attached to the tube wall.
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; she will show you where the thermal cycler is located in E301. Your instructor will start the reaction when everyone's tubes are loaded.
|| 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
|| # of Cycles
| Initial Denaturation
|| 5 min.
| 10 sec
| Final Extension
| 10 min
While the 16S rRNA genes from all of the bacterial species in your soil genomic isolate are being amplified in the thermal cycler, you will have about an hour to work on the soil community profiling assays that you started last week.
However, before you leave today, 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 in Part B. You will also need to set up a gel (described in Part C) 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.
While the amplification of bacterial 16srRNA genes goes on in the thermal cycler, we have about an hour to work on the community profiling analyses that we started last week.
In Lab3 you started some quantitative assessments of your soil communities' ability to digest cellulose, starch, and to solubilize phosphates---all important functional roles. Today while the thermal cycler completes our polymerase chain reactions, you will have about an hour to complete the colony colony counts from the differential media that you inoculated with dilute soil extract last week. We will also talk about how your carbon source utilization profiling is progressing.
Part D: Community Soil Physiological Profiling: EXOENYMES PROTOCOL con't:
Examine the plates for processing of a particular nutrient (starch, cellulose or insoluble phosphates) in each differential culture medium. Remember that these differential media are not selective (they aren't designed to inhibit the growth of any groups of soil microorganisms) but they are Culture-Dependent differential media, in that they allow you to visibly SEE the difference in particular groups of microbes---in our cases, between those that produce and secrete a functional exoenzyme and those that don't. You will count the number of individual colonies showing a clear zone (halo) around the colony (using the plate with 30-300 total colonies) and compare those numbers with the number at the same soil dilution that grew on NA- a general purpose, non-differential medium.
1. Count the total number of colonies on the Nutrient Agar plate and assess total culturable CFUs. Use the soil extract dilution of the plates counted to calculate CFUs/gram of soil (wet weight) for each assessment medium. If you divide the number of colonies counted by the amount of inoculum plated times the dilution factor of that plate, you will obtain the number of cultivatable bacteria per gram of soil.
number CFU/dilution plated*dilution factor = number of CFU/gram
For example, if you counted 150 colonies on the 10-3 plate the calculation is:
150/(0.1ml plated*1X10-3dilution)= 150X104 which in scientific notation is written as 1.5X106 CFU/gram
2. Flood the starch plate with a thin layer of Grams iodine and count the number of colonies that show starch digestion activity as a clear zone or non-blue halo around the colony).
3. Count the number of colonies that show cellulose digestion activity as a clear zone or halo around the colony.
4. Count the number of colonies that show phosphate solubilizing activity as a clear zone or halo.
5. Calculate the % positive for the enzymatic activity for each assay (# positive colonies x dilution factor/total colony count x dilution factor [on nutrient agar] ) X 100. This correction for dilution factor allows you to compare the CFUs counted from different dilutions on plates. If you are able to use control (NA) and test plates from the same dilution (each has between 30-300 colonies), you can omit the dilution factor. This is the total number of CFUs/gram of wet soil of microorganisms able to perform the role of interest.
6. Add your data to the course spreadsheet on the instructor's computer. Be sure to click File Save after you enter your data.
PART E: Isolation of Azotobacter, Hyphomicrobia, Spore Forming, or other interesting Bacteria
Continue to attempt to isolate to pure culture desired groups of bacteria. Directions found in the Protocols section of the wiki at Cuture Media: General Purpose, Selective, Enrichment, Differential, & Assessment of Digestive Exo-Enzymes
Directions for Streaking for Isolation onto new solid media is found at Streaking for Isolation
Your goal is for each student to end up with 3 pure cultures of DIFFERENT genera of bacteria from as many groups as possible.
Once you believe you have pure isolates, continue to subculture to fresh plates each week (isolation streak a colony onto a fresh plate), in subsequent labs you will make a bacterial smear and do a Gram stain and start other tests to explore the physical and metabolic characteristics of this isolate. Generally the medium used is the isolation medium, however, at some point you may want to test the ability of your isolates to grow on nutrient agar. Remember, if you successfully isolated hyphomicrobia your colony should not grow when streaked on nutrient agar. The other cultures may grow as well or better since the nutrient agar we use is rich in nutrients. If your organism grows well on nutrient agar, you can streak on this medium each week and stop using the original isolation medium. Ask you instructor if you are not sure what to do.
1. All culture plates that you are finished with should be discarded in the big orange autoclave bag near the sink next to the instructor table. Ask your instructor whether or not to save stock cultures and plates with organisms that are provided.
2. Culture plates, stocks, etc. that you are not finished with should be labeled on a piece of your your team color tape. Place the labeled cultures in your lab section's designated area in the incubator, the walk-in cold room, or at room temp. in a labeled rack. If you have a stack of plates, wrap a piece of your team color tape around the whole stack.
3. Remove tape from all liquid cultures in glass tubes. Then place the glass tubes with caps in racks by the sink near the instructor's table. Do not discard the contents of the tubes.
4. Glass slides or disposable glass tubes can be discarded in the glass disposal box.
5. Make sure all contaminated, plastic, disposable, serologic pipets and used contaminated micropipet tips are in the small orange autoclave bag sitting in the plastic container on your bench.
6. If you used the microscope, clean the lenses of the microscope with lens paper, being very careful NOT to get oil residue on any of the objectives other than the oil immersion 100x objective. Move the lowest power objective into the locked viewing position, turn off the light source, wind the power cord, and cover the microscope with its dust cover before replacing the microscope in the cabinet.
7. If you used it, rinse your staining tray and leave it upside down on paper towels next to your sink.
8. Turn off the gas and remove the tube from the nozzle. Place your bunsen burner and tube in your large drawer.
9. Place all your equipment (loop, striker, sharpie, etc) including your microfuge rack, your micropipets and your micropipet tips in your small or large drawer.
10. Move your notebook and lab manual so that you can disinfect your bench thoroughly.
11. Take off your lab coat and store it in the blue cabinet with your microscope.
12. Wash your hands.
Write an Introduction section of final paper. Full directions and useful references can be found at Lab 4 Assignment: Assignment: Introduction
This assignment is due at the BEGINNING of Lab 5. Do not come late to lab because you are printing or otherwise completing this assignment and you may NOT work on it during lab. There is a 5% per day late penalty for work for this course and since you have a week or more to complete assignments, illness (unless it is lengthy and serious) does not excuse you from the late penalty.
Continue monitoring and following the appropriate protocols to isolate to pure culture our targeted bacteria.