Lab 3: Continue Soil Microbial Communities & Diversity Project
Today you will continue your identification of the total bacteria in your soil habitat by measuring the concentration of the genomic DNA that you isolated in Lab 2. When your DNA is at the proper concentration (100ng/μL) you will amplify, in a polymerase chain reaction (pcr), part of the 16S rRNA gene from hundreds or thousands of the different species of bacteria (some of which are culturable and most of which are non-culturable) using "universal" bacterial primers and a proof-reading DNA polymerase. If you are unfamiliar with the basics theory behind pcr amplification of target DNA, please review it in at the following web site:
The bacterial primers we will use are called 27 Forward and 1492 Reverse. The numbers refer to positions on the gene where these single strand primers will anneal and where the DNA polymerase will begin to use your soil isolates' genomic DNA as templates to create double stranded sections of the 16s rRNA gene from the various species of soil bacteria in the community. Forward and Reverse refer to the direction the template strand of DNA is copied. As you know, the direction of elongation of new DNA can only proceed in the 5' to 3' direction; therefore, the primer that anneals to base 27 on the 16s rRNA gene template is called the forward primer because DNA synthesis moves downstream on the template. The 1492 Reverse primer anneals to base 1492 on the opposite template strand and DNA synthesis moves upstream from the template. Our primers are called "universal" bacteria primers because they are composed of short sequences of the gene that are common to most Eubacteria (true bacteria). If we were interested in identifying eukaryotic microbes, such as fungi or protozoa, or if we wanted to identify the Archaea (which are neither eukaryotes or true bacteria), we would have to use different primers. Our primer sequences are designed to have sequences that are not found in other non-bacterial ribosomal genes.
Although our primers have sequences of single stranded DNA that are common to the 16s rRNA gene of most true bacteria, the section of the 16s rRNA gene that they will amplify contains highly variable regions of that gene. So variable, in fact, that we should be able to use the DNA sequence of those ~1,500 bases to identify our soil bacteria by comparing the sequences we amplify to the same region of the gene in other bacteria. 16s rRNA gene sequences are archived in an extensive, public database called the Ribosomal DNA Project (RDP) found at | http://rdp.cme.msu.edu/index.jsp;jsessionid=04A058D492CD00120AC01A70AAD7624A .
There are so many 16s rRNA gene sequences archived at this site that we should be able to use their powerful, public search engine to identify our soil bacteria down to the genus or species level, if our pcr, our cloning, and the sequencing of the gene from the clones all works well.
Your careful attention to following the directions exactly in each of the procedures involved in this process will go far to insure the project's success. Keep in mind that the reagents involved are expensive and limited. We will not be able to repeat may of the procedures if you make a signficant mistake. The best way to end up with good data is to read ahead, think about what's happening in each reaction step or in each part of the process, create carefully prepared and organized flow diagrams or outlines in your lab notebook BEFORE you come to lab, ask questions BEFORE you commit yourself if you unsure about a step, and focus on the task at hand.
Today the task is to get the genomic DNA you isolated from your soil sample last week into the proper DNA concentration to amplify the 16s rDNA. Then we will do a "clean-up" procedure to remove the excess dNPTs, primer dimers, etc. from your pcr products. You will stop there for today, but your instructors will use your pcr products to run an agarose gel on a 1/10 volume of them before freezing them away for you. Looking at the purity of your pcr amplification by agarose gel electrophoresis will insure that you have sufficient and pure 16s rDNA to continue the project.
In Lab 4 you will clone the most successful of your pcr amplifications of 16s DNA into a special cloning vector and complete the transformation of those plasmids into gentically modified E. coli bacteria. You will plate the bacteria on selective media so that the bacteria that took up a cloning vector plasmid (containing an antibiotic resistance gene) will grow. Those desirable bacteria are called transformants. The bacteria that did not take up a cloning vector plasmid will not have the antibiotic resistance gene (non-transformed bacteria). The non-transformants should not be able to form colonies on the media with the antibiotic.
We hope that we have hundreds of well isolated colonies from the transformation and selection process. In Lab 5, each pair of you will choose 96 if those cloned bacteria, each containing an inserted 16S rRNA gene from (we hope) many different bacterial species from your soil samples, and get them ready to send away, as glycerol stocks, to a company that will purify these plasmids from out E. coli and then sequence the inserts. The sequencing facility is able to use conserved primer sites on our vector to be able to sequence the inserted 16S rRNA genes. We should get our sequencing data back within two weeks. At that point, after having a tutorial session on how to analyze the sequencing data using the RDB public database, we will be able to identify many of the bacteria in our soil habitat. We hope those identifications will include a wide variety of the flora that we did not culture; however, since we will randomly pick identical looking transformed E. coli clones and those clones randomly took up vector plasmids that randomly inserted an amplified 16s rRNA gene from genomic soil DNA isolate, we have no control over how representative our data will be of the soil bacterial community in your habitat.
The best approach, if we could afford it, would be to use 454 pyrosequencing to really sample the PCR amplicons from our soil community. Pyrosequencing is a relative new method of generating about 30,000 reads per sample rather than the 96 that we will get from our Sanger sequencing approach. The following links give more information comparing the two sequencing processes: | Link to Roche website with animation explaining 454 pyrosequencing process;
Link to other information on 454 pyrosequencing compared to traditional Sanger sequencing: Wikipedia comparison of 454 and Sanger sequencing: ;
Link to a PNAS 2006 paper comparing the two methods:| http://www.pnas.org/content/103/30/11240.full
Unfortunately, to do 454 pyrosequencing on our soil samples, it would cost about $10,000; that's well over the course budget! Despite our less that optimal approach to identifying the diversity of bacteria in your soil community, you should get a good sense of what's there. In fact, the silver lining to using our limited approach is that you might not feel entirely positive about trying to analyze 30,000 reads from pyrosequencing for your final paper.
Part A:Measuring the Concentration of DNA
Your instructor has done Part A, measuring the DNA concentration of your soil, for you and sent the result to you by email. You can read about how this measurement was accomplished in the paragraphs below. We will not need to dilute our DNA and we have decided not to try to concentrate it (for fear of losing our little DNA pellet in the process) so you can skip directly to Part B, setting up your 16S rRNA gene amplification by polymerase chain reaction.
There are two Nanodroppers in the BISC Equipment room, L308. Both the ThermoScientific NanoDrop 2000 and The NanoDrop ND-1000 Spectrophotometers measure DNA by taking Absorbance at A260nm. These spectrophotometers use only 1 microliter of sample and do not require cuvettes. The sample is held in place by fiber optic technology and surface tension that holds the sample in place between two optical surfaces that define the pathlength vertically and dynamically. Measurement can be assessed in a range of 2 to 3700nm/microliter dsDNA. These are expensive machines so make sure you follow the directions carefully and ask your instructor for guidance as needed.
More information is available from the manufacturer's website at: | http://www.nanodrop.com/HowItWorks.aspx
Using the Nanodroppers
1. Clean the upper and lower optical surfaces of the sample retension device by pipetting 2 microliters of clean deionized water onto the lower optical surface. Close the lever arm and tap it a few times to bathe the upper optical surface. Lift the lever arm and wipe off both optical surfaces with a Kimwipe.
2. Open the NanoDrop software from the Desktop of the computer and select the nucleic acids module.
3. Initialize the machine by placing 1 microliter of clean deionized water onto the lower optic surface, lower the lever arm, and select initialize from the NanoDrop software. Once initialization is complete (~10sec.), clean both optical surfaces with a Kimwipe.
4. Perform a blank measurement by loading 1 microliter of Solution 6 (10mM Tris) and select Blank. Note that this blanking step may use something other than Tris depending on what your sample is dissolve in. Often the blank will be deionized water if you have concentrated your DNA sample already with the ethanol precipation and resolubilized it in water.
Note that as in traditional spectroscopy, the blank will be subtracted from subsequent measurements. If you want to determine the contribution of a specific buffer or diluent, measure the buffer first using distilled water as a blank. If the buffer does not contribute to the A 260nm reading, then deionized water will be fine to use as the blank. The water or buffer should always be measured to be sure that the instrument has been zeroed properly. The measurement of water or buffer should be zero or very close. All measurements are automatically normalized to 340nm.
5. Measure the nucleic acid sample by loading 1microliter of sample and selecting "measure". Record your DNA concentration. Once the measurement is complete. Clean both optical surfaces with a Kimwipe and the machine is ready for the next sample.
You should ensure that the appropriate constant (50 for dsDNA or 40 for RNA) has been chosen. The software automatically calculates the nucleic acid concentration. If the calculation is done by hand, the A260nm is represented as a 1cm path for convenience, even though 1-nm and 0.2nm paths are actually used during the measurement cycle.
When the last sample was been measured, clean the sampling device by repeating step 1.
To Dilute or concentrate your DNA for pcr amplification of 16S rDNA
The final volume of eluted DNA from the DNA isolation procedure should be ~100μL. We would like for our DNA to be at a concentration of ~ 100ng/μL for our pcr amplification. If you didn't achieve that concentration, we will adjust our pcr reaction template volume as described in the PCR protocol rather than using the ethanol precipitation concentration procedure described below. The concentration protocol is described here for completeness, but we won't have time to do it today.
Diluting the DNA
If your isolate is more concentrated than 100ng/μL, you can dilute it in sterile water or 10mM Tris (solution 6). Ask your instructor which is preferred. To calculate how to dilute your DNA, you will use the formula V1 x C1 = V2 X C2, where V1 is ~100μL volume of your eluted DNA from the PowerSoil isolation, C1 is the DNA concentration you obtained from the Nanodroper, C2 is 100ng/μL (the conc. you desire), and V2 is what you will solve for. Please check with your instructor after you have made this calculation and BEFORE you dilute your DNA.
Concentrating the DNA:
Ask your instructor if you should concentrate your DNA through the ethanol precipitation procedure described next, OR just adjust the volume of template DNA to use in the pcr reaction described in part B. (The danger of doing the ethanol precipitation with small amounts of DNA is losing the tiny DNA pellet and ending up with nothing.)
The DNA may be concentrated by adding 4 microliters of 5 M NaCl to your ~100μL DNA isolate and inverting 3-5 times to mix. Next, add 200μL of 100% cold ethanol and invert 3-5 times to mix.
Line up the hinge on the cap of the microfuge tube so that it points out before you centrifuge at 10,000 x g for 5 minutes at room temperature. By placing the hinge out, you will know where your DNA pellet should be in the tube after it's centrifuged. Sometimes these pellets are almost invisible and you don't want to suck it up and discard it when you decant the supernatant in the next step. Look hard for the tiny white DNA pellet after centrifugation and keep your pipet tip away from it when you decant ALL the liquid using your P200. The pellet must be completely dry and every trace of ethanol must be removed before you try to resolubilize the DNA in water. Remove residual ethanol in the speed vac in E301. Ask your instructor to help you find and use it. While your sample is in the speed vac for 5 minutes, calculate the volume of water you will need to resolubilize your DNA to the appropriate concentration. Check your calculations with your instructor BEFORE you add any water to your dried pellet of DNA. Use the next paragraph for how to calculate the dilution.
Use V1 x C1 = V2 x C2 to determine how much water you will eventually add to your dry DNA pellet. Remember that you know the original concentration, C1, from the nanodrop measurement and you know the original volume, V1. You also know C2, 100ng/μL, the concentration you desire. You want to solve for V2, which is how much volume you desire to solubilize your pelleted DNA in to give you the desired concentration of 100ng/μL.
Be sure that you ask your instructor to check your calculation before you go on the next step.
Resolubilize your precipitated DNA in the calculated volume of sterile water. After adding the appropriate volume of diluent to your tube, vortex vigorously at highest speed for 2 full minutes to make sure your DNA is completely in solution.
When your DNA is at the appropriate concentration, 100 nanograms/μL,
proceed to the next step: Amplification of 16s rDNA by Polymerase Chain Reaction.
Amplification of Bacterial Genomic DNA by Polymerase Chain Reaction to ID Soil Unculturable Flora
Our genomic DNA isolation has, no doubt, resulted in a mixed DNA population from a myriad of microorganisms as well as, probably, some contaminant DNA from plants, insects, or other life forms in the soil community. Since we are only interested in the scope of our bacterial population in this study, 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 amplification from the template DNA that begins with this common region, but ends, after 35 cycles of polymerase chain reaction in a thermal cycler, with a pcr product containing hundreds of unique copies of 16s rDNA, allowing identification of much of the bacterial flora present in the soil community, most of which is unculturable by conventional techniques.
Part B: 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 extremophilic, Thermus acquaticus living and reproducing in boiling hot springs. We are using a different polymerase, Finnzyme's Phusion High-Fidelity Polymerase, a proprietary reagent that uses a novel
Pyrococcus-like enzyme with a processivity-enhancing domain. Phusion DNA Polymerase generates long templates with an 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 4μL of 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, 30μL DNAase free water, 2.5μL of each of 2 primers).
If your soil isolate DNA concentration was less than 20ng/μL, you will add 5 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 5) and enough DNAase free water so that the number of microliters of DNA + microliters of water =5. Example: Your DNA conc. was 33ng/μL. 100/33 = 3.3 so you would add 3.3μL of DNA and 1.7μL of DNAase free water. Since your pcr tube already has 10μL master mix, 30μL DNAase free water, and 2.5μL of each of 2 primers, the total reaction volume for everyone will be 50μL.
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.
| Component || amt. in a 50 μl|
| Final Conc.
| 30μL already in tube.|
Want to achieve
total of 50 μl reaction vol.
Add from 0 - 4μl
| 2x Phusion Master Mix
|| 10 μl
| 27F primer
|| 0.5 μMolar
| 1492R primer
|| 0.5 μMolar
| template DNA
|| 1-5 μl
|| optimum is 100ng of DNA/reaction
The cycling program is shown below. 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 proceed with the other parts of the lab.
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 in Part C.
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 |
Part C: Clean Up of pcr product using Epoch BIoLabs GenCatch PCR CleanUp Kit
Notes before Starting:
Make sure 95% ethanol has been added to Buffer WS before first time use (see bottle label for volume).
All centrifuge steps are carried out at 17,900 x g (13,000) rpm) in a conventional tabletop microcentrifuge at roomtemperature.
1. Measure 500 μl of Buffer PX using your P1000 and add part of it to your thawed 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 no more than 700 μL of the pcr product/bufferPX mixture created in step 1 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 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. If you applied all the pcrproduct to the spin column in step 3, skip this step and proceed to step 6.
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 column by applying 700 μL of Buffer WS to the spin column. Note that WS Buffer is different than the buffer used in step 6. Centrifuge the column for an additional 1 min. Discard the flow through
9. Centrifuge the spin column in the same collection tube at full speed for 3 more minutes to remove 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 50 μl 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).
IMPORTANT: Ensure that the elution buffer 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, and 28 μl from 30 μl elution buffer.
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
Agarose Gel Electrophoresis of Cleaned PCR Products
To see if you successfully amplified the 16s rRNA gene and not anything else, your instructor will "run a gel" on your cleaned pcr products. To run a gel means that she 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. She will 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?
Procedure for Agarose Gel Electrophoresis of PCR products
Load 1/10 of the total volume of pcr product (1 microliter minimum)
Mix 1 volume of Loading Dye with
5 volumes of purified DNA. (Mix the solution by pipetting up and down before
loading the gel.)
Loading dye contains 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. We will use bromophenol blue this time.
Give your cleaned up pcr product to your instructor to freeze after the gel is run, making sure your pcr product is clearly labeled with your team, lab section, soil identifier code, date. In the next lab you will clone the fragments of 16s rDNA from the soil bacteria flora that are in your pcr product into a special genetically engineered cloning vector. That vector will be transformed into competent genetically engineered E. coli bacteria and they will be plated on selective media to find cells containing the 16s rDNA insert that we can send away for sequence analysis to determine the identity of some of the bacterial flora in your original soil sample.
Isolation & Characterization of Cultured Bacteria from a Soil Habitat
Remember that you are working on two aspects of our Soil Diversity Project simultaneously: We are getting a sense of the total bacterial community through sequencing of the 16S rRNA genes (rDNA) from the genomic DNA isolation of our soil sample and we are also using traditional culture techniques AND DNA sequencing to isolate and characterize a few of the interesting culturable bacteria. You began the isolation and selection of some culturable bacteria of specific groups from your soil sample through the use of selective and enrichment media in Lab 2. You also started a standard plate count of the culturable organisms. Today you will complete the plate count to get one kind of enumeration of the microbes in your soil community. You will also continue the process of isolating and selection of specific groups of microbes based upon unique or distinctive metabolic, morphologic, or behavioral characteristics.
Part D: Finishing the Standard Plate Count of Culturable Soil Microbial Flora & Observing Colony Morphology in the Streak Plates
Activity D-1: Find a plate that contains 30-300 colonies (there should be only one if you did your 10 fold serial dilution correctly). Count all the surface and subsurface colonies on that plate. The colonies can be more easily counted by using a Quebec Colony Counter which allows proper illumination, a grid overlay and by slight magnfication of the plate surface. There are two in the lab.
Calculating the number of bacteria per gram of soil
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. If it is clear that a culture plate has well over 300 colonies or under 30, designate it as "invalid" in your lab notebook.
CFU = number counted on plate/(diluent plated*dilution of plate counted)
Things to Consider:
Why did we perform so many dilutions?
Why should you only have one plate with 30-300 colonies?
Don't forget that these plates are also a potential source for isolates, particularly those in the aerobic spore forming group and Actinomycetes (which includes the Streptomycetes).
Activity D-2: Observe your streak plates. Is there a great deal of growth in zone 1 of your streak plates? How good was your technique at streaking for isolation? Did you get well isolated colonies by the 3rd or 4th section? If not, consult with your instructor and ask her to watch your technique.
Activity D-3: Examine the colonies on the dilute and full strength nutrient agar steak plates. Record general observations in your lab notebook about the number and variety of colonies or the growth observed. Draw and/or take pictures of some of the most numerous and some of the most interesting colonies. Compare the appearance of microorganisms on general purpose media to the colonies growing on your selection/enrichment media.
Observing different colony types, based on colony size, shape, elevation, color, and consistency.
Colonial characteristics are best observed where individual colonies are well isolated. Scan your plates to find the different categories described below. Note differences in numbers and types of colonies in different media. We wish to avoid working with fungal colonies, which generally appear as "fuzzy" growth.
Size - pinpoint, small, medium, large, very large, and spreading.
Color - lemon yellow, golden, creamy, gray and translucent, white
and opaque, red, pink and so on.
Appearance - shiny or dull
Shape -round, irregular, and so forth.
Elevation - flattened, heaped up, concave
Consistency - dry and friable, easily broken up, sticky and stringy,
and so on.
Odors - try to describe them.
Activity D-4: You will continue and/or start the process of isolating pure colonies from your various enrichment protocols.
While unlikely, it is possible that a few fast growing bacteria in the care of avid microbiology students may already be at the pure culture stage (have been isolation streaked at least 2 times and all the colonies growing from the last streak look identical). If so, you should follow the directions in LAB 4 Activity 4C-1 to make stock cultures before you do any tests on your isolates. If you have time today and enough of the pure colony left after inoculating your stock slants, you can perform Gram stains on the bacterial colonies that you think are in pure culture, following the directions in LAB 1, Making a Bacterial Smear and GRAM STAIN.
Most students, however, will probably not have bacteria in pure culture by this week. In this case you will begin or continue the isolation of interesting bacterial colonies from your enrichment or general purpose media. Select several different looking colonies from each of the plates (or other forms of culture) worked on over the past week. Remember that our goal is for you and your partner to isolate and identify as wide a variety as possible of interesting organisms, so please chose colonies that are different in appearance from each other and that come from as many as possible of the selection/enrichment media.
Please streak each selected colony for isolation on a new plate of the same media where the colony was found. After selecting a colony on a plate DO NOT touch your loop to any part of the plate other than directly on the colony of interest. There may be other types of bacteria living on the media that have not reproduced enough to form visible colonies. DO NOT select any colony that is likely to be a fungus. Check with your instructor if you are unsure about how to identify fungal colonies. Keep in mind that you will not necessarily continue the identification of all the colonies you select this week. Some isolates may die off and others may not provide enough variety to your final collection, so select as many different looking colonies as possible this week.
Continue with all the enrichment protocols you have in progress. Each student will probably be at different steps or stages in the more time consuming enrichments you began last week.
Things to Consider When Using Selective or Enrichment Media"
- What are the ingredients in these "enrichment" media or protocols that target a desired group's growth restrictions or preferences (nutritional needs, concentration of oxygen, light, pH, temperature)?
- What are sources of C,N,S, P in the various enrichment media?
Your eventual goal is to use DNA sequencing, morphological, metabolic, and biochemical tests to identify some bacteria in your chosen soil community and to examine the role(s) these bacteria play in the soil; but, first, you need to be sure they will grow successfully and are in pure culture.
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.
Create a Plan to Isolate, ID, and Determine the Role(s) of a Diverse Group of Bacteria in a Soil Community
Get together with the other members of your soil sampling team and devise a plan for ending up with isolates of at least 16 (4 each) unique bacterial genera found in the soil community you sampled. In your attempt to end up with at least one from each of the 11 groups described in the Enriching for Soil Bacteria in a Mixed Population, you and your partners should not limit yourself to 4 colonies yet. It is highly likely that some promising bacteria will not survive to the end of the identification or that several of the same species of bacteria are growing on multiple media and may look a bit different on each type. Work with your team to assign different bacterial groups to each member. Each of you should continue to isolate and characterize at least two or more different looking colonies from the groups that are not ready for isolation streaking yet and that are likely members of your assigned groups (8 probably different bacterial species from a minimum 4 different groups).
Keeping track of the highly variable protocols for selective enrichment is challenging. Your best organizational skills are required. You will be expected to come in at various times, on your own, to start, continue or complete an isolation or characterization test. We will make every attempt to make the media and reagents that you require available when you need them, but since everyone is proceeding with different tests and media at different times, that availability requires advance planning on your part and communicating with your lab instructors your needs or desires well in advance of time of use. Remember that because this is an investigative, project based lab course, your instructors do not know the identity of the bacteria you are culturing from your soil habitats. The success of this project is in your hands. Early and continual updating of the plan you devise is crucial. You will not turn in this plan for a grade next week, but your instructor would like for the team to create a master plan with a preliminary time line and check it with your instructor.
Write a Materials and Methods Section
Some of the things we will stress in this lab, other than acquiring experience with the tools and techniques of the microbiologist, are analysis and presentation of experimental data and scientific writing. Our semester long project will end with a group presentation and an individually written final paper. The presentation will model a poster presentation given to community of peers at a scientific conference and the final paper will be written in the form of primary research report. The topic of both is Bacterial Diversity in a Soil Community: Roles and Relationships Among Soil Flora.
It can be a daunting task to write a single paper on several months of experiments, particularly when those experiments result in a huge amount of disparate data. To help you manage your project more effectively, and to prevent you from having an overwhelming amount of analysis and writing to start from scratch at the end of the semester, we have designed assignments throughout the semester that will be evaluated and graded. The feedback you receive on parts of the data analysis or on drafts of typically problematic sections of the paper, we hope, will help you work on the paper throughout the semester and help make the final submission exemplary.
Your first assignment in this process is to write up the protocols completed, so far, in Identification of the Bacterial Soil Community by 16s rRNA Gene Sequencing. That would include: acquiring a soil sample from a particular habitat, isolating genomic DNA from it, amplifying 16s rDNA by polymerase chain reaction (this amplification includes the adjustment of DNA concentration, the pcr product "cleanup" and the assessment of success by agarose gel electrophoresis). Although those procedures take up substantial space in the lab wiki, turning all those protocols and descriptions into a Materials and Methods section should require a page or less of text (double or 1.5 spaced). Remember that the Materials and Methods section of scientific paper does NOT read like protocol descriptions in the wiki. The emphasis is NOT on what you did in lab, step by step, in a time course description of your lab days, but rather on a BRIEF description of the process of turningstarting material into whatever the goal material may be. In this case, the starting material is a particular soil sample and the goal is to end up with lots of double stranded DNA segments of a particular part of the 16s rRNA gene from as many of the bacteria found in your soil community as possible.
Each part of Methods & Methods in your final paper will be a separately titled section. The section titles should emphasize the experimental goal of that part of the larger experiment, rather than being a more general descripiton of procedures or equipment. Therefore, a good title for this Methods section would stress the amplification of _____ from________ by____, rather than "Pcr of_____. There may be titled sub-sections, or not, in each Materials and Methods section. Remember that the goal of a Methods description is to give a future investigator enough information to accomplish the same experimental goal, but, perhaps in a different context, scale, or using a different starting material. Therefore, you need to include all the essential information so that such a future investigator can make or buy all reagents and complete the experiment successfully to its goal, but you do not have to include details that an experienced investigator would infer (such as specifying that you "mix" after combining reagents or giving the type of general equipment such as the tube type you used). NEVER, EVER use the word "tube" in Materials in Methods. We often tell you, in a protocol, to do ____ to the "tubes". In the context of a lab manual procedure description, that's fine. For Materials & Methods, you must be more specific. It's what's happening to the materials in the tubes that is important. Tubes and plates are just pieces of plastic. Your methods section should not read like a series of "did this, then did that" descriptions, but should, instead, describe how you acquired and processed the starting material to end up with the ending material.
Please look at the Methods sections of published scientific research papers (there are many in the References folder in your First Class lab conference) to see how the protocols in this wiki differ from Materials and Methods descriptions in a scientific paper. Notice that M&M can be very short, particularly if you have used a kit or a previously published protocol for a part or all of the experiment. If some part of your work is published elsewhere, such as in a journal article's methods section or in a kit manufacturer's website, you may reference the url of the kit manufacturer or use a journal article citation to the previously published directions. Remember, however, that in the body of the paper, such a journal citation in the style of the journal Cell is given as (First Author's last name/ Year of publication), but there must be a FULL citation of the article in the References list at the end of the paper. If you use such a short-cut (and it is fine or desirable to do so), include a Reference page with the full citation as part of this assignment so you won't have to search for it later.
Since most of our protocols come directly the manufacturer of the main reagent or kit and others come from well-established published sources, this assignment should not be extensive in length. To write about how you accomplished your experimental goals succinctly and clearly, you must thoroughly understand them. It does not require thorough understand to follow the directions in the wiki and end up with a successful experiment; however, to write about your experiments in such a way that someone who is not part of this course can understand what you did and how it contributed to your overall goal... that's is not so easy. If you make sure that everytime you leave lab you understand how each day's work fits into the project's overall goals, writing M&M will be easily manageable. Don't wait until the end of the semester to figure it all out. If you want to keep practicing converting our protocols to M&M as you complete them rather than limiting yourself to those we ask you to turn in for graded assignments, you will make the writing of your final paper, much easier.
Please do NOT reference the wiki, as it not a primary source. Keep in mind that even though you can ease the burden and length of this assignment greatly by referencing other journal articles or a reagent manufacturer's web site (such as the New England BIolab's website directions for using Phusion® High-Fidelity PCR Master Mix with HF Buffer-- the reagent we used to amplify our target DNA by pcr (found at: | http://www.neb.com/nebecomm/ManualFiles/manualF-531.pdf), the manufacturer does not include the sequences of the primers we used (since those are unique to your target DNA) nor the exact thermal cycler program (since that is tweaked for the length of the DNA section you want to copy and for the relative CG content of the template DNA). Therefore, you must give more than a brief citation and web address for most experimental descriptions. In the case of the pcr amplification using a commercial master mix, you must be specific about the template DNA, the primer information for amplifying 16s rDNA, and give the exact thermal cycler program you used.
There is an extensive handout on writing a Methods section in the Resources section of this wiki. In this course we will use the citation style of the journal Cell, one of many scientific citation styles. The Wellesley Library has electronic access to Cell, so you can look at recent issues to see how the authors of research reports formatted their citations and use those Reference pages as models when formatting your references. Attention to detail is crucial in scientific writing. You must follow the citation style requested exactly and uniformly.
This assignment is due at the BEGINNING of Lab 4. 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 enrich and isolate the culturable bacteria.
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