BISC209/S11: Lab6

LAB 6: Finishing the Culture-Independent Identification of Bacteria by 16s rRNA Gene Sequence Analysis

To summarize our culture-independent work, you have extracted genomic DNA from a soil sample, amplified the 16S rDNA by pcr using "universal bacterial primers", inserting the different 16s rDNA fragments from your pcr product into a cloning vector, transformed special genetically modified E. coli with a plasmid in order to separate 16s rRNA genes from different bacterial members of your soil community. Now you are ready to pick transformants that successfully incorporated the plasmid vector genes.

Activity::

Preparing your clones to send away for sequencing analysis of your 16S rRNA gene
When you examine your transformation plates after their initial overnight incubation, there should have been hundreds of well isolated colonies. In theory, each of them should contain the vector plasmid with an insert of the 16s rRNA gene from one of your soil sample bacteria. Since the vector plasmid contains a kanamycin resistance gene, kanamycin resistance is a selectable marker. The genetically engineered strain of E. coli that we transformed is sensitive to kanamycin UNLESS it is expressing the kanamycin resistance gene on the plasmid. E. coli that did not take up a cloning vector plasmid and express its genes do not form colonies on media with kanamycin. Kanamycin is an antimicrobial compound that disrupts bacterial protein synthesis and kills the cells.

We know each of the vector plasmids in the transformed E. coli growing on the plate contains a 16S rRNA gene insert from the genomic DNA isolated from your soil sample for two reasons. First, there is a ccdB gene in the insertion region of the vector plasmid. That gene, ccdB, means "control of cell death". That gene, when not disrupted, expresses the ccdB protein that poisons bacterial DNA gyrase, causing degradation of the host chromosome and cell death. But the presence of your 16S rRNA gene insert has disrupted the ccdB gene and turned off the protein that inhibits DNA gyrase, allowing the cell to live, replicate and form a colony that should appear white, NOT blue. The second reason that we know the white colonies are transformed with the vector plasmid and that the plasmid contains our insert is that there is a lacZ gene positioned next to the ccdB gene in the insert area and when it is disrupted by insertion of your 16s rRNA gene, it turns off expression of the lacZ gene product, beta-galactosidase. Beta-gal is in enzyme that catalyzes the breakdown of several substrates, including lactose and X-gal. X-gal is a colorless substrate that is is cleaved into a blue colored product by beta-galactosidase. Your Luria-Bertoni agar medium contains both kanamycin and Xgal. If you saw blue colonies, those bacteria are daughter cells from a vector transformed E. coli, BUT the vector plasmid probably does not contain the 16s DNA insert we seek. Therefore, you only want to pick "not-blue" colonies to send away for sequencing of the insert. We hope that there are hundreds of these not-blue colonies on your plate (but not so many that they are not well separated from each other). Our goal is to find 16s rRNA gene fragments from DIFFERENT soil bacteria in many transformed clones, but we have no way of detecting right now which colonies contain a 16s rRNA gene from different soil bacterial species because all will be identical looking non-blue colonies on these plates.

Each lab section will be allowed to fill (3) 96 well sterile blocks with their clones. Follow the directions below, carefully, to inoculate each well with a different, well isolated non-blue colony.

Preparing Over-night Cultures to send away for 16s rRNA gene sequencing

1. We need to keep track of which DNA sequences come from which sampling site. Therefore, you should make sure that each sampling site has a separate, well-labeled 96 well plate. Start adding clones at the beginning (wells A1-A2, etc). Be sure the plate is labeled Tues. or Wed. lab, and that it has your soil sample identifier code for the wells used. If you have extra wells (not enough non-blue clones), let your instructor know.

2. In the hoods in the lab, you will find 3 prep areas for transferring colonies to the 96 well block.
You will use your P1000 to inoculate 1 ml (1000μL) of LB broth with 50 μg/ml kanamycin (NO X-gal) into each well of your block. We suggest you inoculate one well at a time. First add the medium then add your colony.
3. Find and select well-spaced, white, colonies on your transformation plates.
Use the flat end of a sterile toothpick to pick up a single colony. Be careful NOT to touch any of the area of the plate around the colony with your toothpick! Place the toothpick in a singe well of your 96 well block. Leave the toothpick in the well as added insurance that will know which wells have been inoculated!!!!
4. Once all the wells assigned to you and your partner are filled with toothpicks, carefully pull out each toothpick by wiping it on the edge of the well (to scrape off the organism) on a side of the well that will allow you to discard the toothpick without the chance of dripping this well's contents into another well. BE CAREFUL not to cross-contaminate any wells!!! Discard the toothpicks in the autoclave bag.
Once the block is completely full, apply the sterile sealing mat carefully and label the plate. This label should include your lab section day, team colors, sampling site codes, and the date. Make sure that this identifying information is also on the template. Place full blocks carefully on the platform shaker and tape them down before turning on the shaker. Your block will incubate with constant shaking at 37C overnight. Give your instructor the completed template.

Preparing Glycerol Stocks from your Overnight Cultures(Your instructor will do this part for you so make sure that your plate and wells are clearly identified!)

1. Pipet 50 μL of 50% glycerol in each well a 96 well Costar round bottom plate.
2. Mix each overnight culture from the 96 well block by pipetting up and down and transfer 50 μL of each culture into a separate well of Costar plate. Mix well.
3. Seal the plate with an aluminum foil special seal and label the plate clearly: Wellesley College, BISC209, Tues or Wed (for lab day), (for sampling site codes) and the date.
4. Freeze at -80C and send away for sequencing on dry ice.
The sequences should come back in a week or two.

Culture-Dependent Analyses

Assessing Bacterial Morphology and Characteristic Arrangement and Cell Wall Composition by Gram Stain

Use 1/8 of a well-isolated colony of each bacterial soil isolate to make a smear slide, Smear Slide Preparation, and to perform a Stains: Gram Stain.

Activity: Preparing a bacterial smear slide
1. Label two clean, glass slides with a graphite pencil on the far left of the slide with the identity of your three isolates on one slide and and the identity of two control cultures on the other slide SE (Staph. epidermidis), SM (Serratia marcescens), Mix. (The decolorizer in the Gram stain can remove your labels if you use pen or wax pencil.) By convention, labels (top to bottom) match smears (left to right).
2. Place three very small loopfuls of deionized water on each slide as far from each other as possible. (You can use the deionized water bottle on your bench; remove the cover and dip your loop in since sterility is not required for this step.
3. Flame the loop, allow it to cool for a few seconds and touch the cooled loop to a colony of S. epidermidis , picking up a TINY bit of white growth from the bacterial colony. An invisible amount of growth obtained from just touching the cooled loop to the colony is fine.
4. Place the loop with the bacterial growth into the drop of water on far left of the slide. Use a circular motion to make a smooth suspension of the bacteria in the water. Stop when there is a circle of emulsified bacteria about the size of a nickle on the slide. Be sure to leave room for the adjacent drop of water to be spread to a similar size without mixing the two smears.
5. Reflame the loop.
6. Repeat step 4 with the Serratia marcescens in the middle drop of water and then, without flaming your loop, touch the loop to the drop of water on the far right and mix briefly.
7. Reflame your loop and touch it to a Staphylococcus colony again. Place the loop in the far right drop of water mixing it with the Serratia and spread the drop as in step 4 to create a mixed smear.
6. Allow the slide to air dry completely! Be sure all the water on the slide has evaporated before proceeding to heat fixation!!! This drying step is crucially important. If you are impatient, you will "explode" the cells in the next step .
7. Heat fix (to kill and attach organisms to the slide) by quickly passing the slide (smear side up) through a flame 3 times. Use a clothes pin or slide holder and avoid contact with hot glass.

An example of a multiple smear labeled slide:

The Gram Stain

Background on Using Stains in Bacteriology
The first of the dyes most useful to bacteriologists was a reddish violet dye, mauvein, synthesized in England by William. H. Perkin, and patented by him in 1856. This synthetic dye and others were immediately appreciated by histologists, but were not applied to bacterial cells until Carl Weigert (a cousin of Paul Ehrlich) used methyl violet to stain cocci in preparations of diseased tissue in 1875. Subsequently, the use of various synthetic dyes for bacteriological preparations developed rapidly when they were promoted through the publications of Robert Koch and Paul Ehrlich.

The synthetic dyes are classified as acid dyes, or basic dyes, depending on whether the molecule is a cation or an anion. The introduction of the terms acidic and basic was unfortunate because it would be more revealing to refer to them as cationic or anionic dyes. A look at the structural formula reveals the nature of the dye.

Each dye molecule has at least two functional chemical groupings. The auxochrome ionizes and gives the molecule the ability to react with the substrate, while the unsaturated chromophore absorbs specific wavelengths of light. The color of the solution obtained is that of the unabsorbed (transmitted) light. To be a dye, the molecule must have both auxochrome and chromophore groups. The auxochrome is usually an ionized carboxyl, hydroxyl, or pentavalent nitrogen group. The chromophore may have unsaturated nitrogen bonds such as azo (-N=N-) indamine (-N=), nitroso (-N=O) or nitro (O-N=O), groups; or unsaturated carbon to carbon, carbon to oxygen, or carbon to sulfur bonds, such as ethenyl (C=C), carbonyl (C=O), C=S, or the quinoid ring (= = =).

Resonance is also important to color. In crystal violet, an electron resonates between the three benzene rings. As the pH of the solution is lowered, the resonance becomes more and more restricted. When the resonance is restricted from three to only two benzene rings, the solution turns from violet to green, and then to red when resonance between the two rings ceases.

Cationic dyes will react with substrate groups that ionize to produce a negative charge, such as carboxyl, phenolic, or sulfhydryl groups. Anionic dyes will react with substrate groups which ionize to produce positive charges, such as the ammonium ion. Any substrate with such ionized groups should have an ability to combine with cationic or anionic dyes. Generally, the most important staining substrates in bacterial cells are proteins, especially the cytoplasmic proteins; however, other substances also have dye affinity. These include amino sugars, organic acids, nucleic acids, and certain polysaccharides.

Sudan III, or sudan black B, is a popular stain for fatty material. It does not have an auxochrome group, and is insoluble in water, but soluble in fatty material. When a solution of sudan black B in ethylene glycol is placed over bacterial cells, the fatty material will dissolve some of the dye and thus take on the color of the sudan black. The staining effect is purely a solubility phenomenon, and not a chemical reaction, or physical adsorption.

There are many stains that can reveal the morphology of the cell, and some simple stains, such as methylene blue, are quite good for viewing bacteria. The Gram stain is especially useful because it not only reveals bacterial morphology, but also is a differential stain. A differential stain differentiates organisms. (A differential stain shows a visible difference between different groups of organisms based on some characteristic they do not share, even though the procedure to stain the different looking organisms is the same). The Gram stain relies on cell wall differences between groups of bacteria.

The Gram staining procedure as it is done today, involves: a) primary staining of all cells with crystal violet, b) precipitating the primary stain dye within the cells with iodine (a mordant), c) removing the dye-iodine precipitate from some cells (the Gram-negative) with a decolorizer such as 95% ethanol, acetone, or n-propyl alcohol, and d) counter-staining of the decolorized cells with safranin. Organisms that retain the crystal violet primary dye are termed Gram-positive, while those which lose the primary stain and show the red safranin counter-stain are termed Gram-negative. This differentiation is not absolute, because it is based on the differences in the rate at which the primary dye is lost from the cells. If you over decolorize for too long or with too harsh a decolorizer, Gram-positive organisms will appear Gram-negative. Truly Gram-positive cells, such as Bacillus subtilis or Staphylococcus aureus, will not retain the primary dye if the iodine step is omitted. Criteria for a true Gram-positive state include the requirement of iodine following the crystal violet.

Since the term Gram positive or Gram negative actually refers to a type of cell wall, not all organisms that retain the primary dye of the Gram procedure are really Gram-positive, because they lack that particular cell wall composition. For example, Mycobacterium species have a different type of cell wall but they will take up and retain crystal violet if you use heat. In this case, the crystal violet will be resistant to harsh decolorization and be retained. However, a Mycobacterium type cell wall does not require the use of a mordant, like iodine, to precipitate the stain. True Gram-positive organisms do not retain the primary crystal violet without precipitation by a mordant. Using crystal violet with heat, harsh decolorization, and no mordant describes an acid-fast stain rather than a Gram stain. Mycobacteria have an acid-fast type cell wall; they are neither Gram-positive or Gram-negative, despite the fact that they will appear purple if you do a Gram-stain on them.

Activity: Preparing a Gram Stain

The Gram stain is a standard staining technique useful for the identification of culturable bacterial organisms and you will perform it now.
Use the slide prepared in Activity 2 and follow the Gram Stain Protocol found below and in BISC209: The Gram Stain in the protocol section of this wiki.

Gram Stain Procedure:

To Gram stain the bacterial smear slides, do the staining protocol from start to finish on one slide at a time. You must be careful to apply the staining reagents liberally so all the smears are evenly and completely covered and you must be sure to expose each smear to each reagent for the same amount of time.
1. Place your smear on the staining tray. It is important that the slide be level during staining so use paper towels under the tray to get it leveled. If you do, it is much easier to be sure that your smears will be covered evenly with each reagent.

2. Dispense just enough Crystal Violet solution (0.5% crystal violet, 12% ethanol, 0.1% phenol) to completely cover each smear and stain for 1 minute. (Crystal violet is the primary stain.)

3. Rinse the slide by lifting it at a 45 degree angle (using gloves or a clothes pin or slide holder) and use a squirt bottle to direct a very gentle stream of water slightly above the top smear. Rinse until the waste water coming off at the bottom is relatively clear; drain off excess water by touching the edge of the slide to a paper towel.

4. Dispense just enough Gram's Iodine (mordant)to completely cover each smear. Let stand for 1 minute. Rinse thoroughly with a gentle stream of water as in Step 1.

5. Lift the slide at a 45 degree angle and drip Decolorizing Reagent (80% isopropyl alcohol, 20% acetone) down the length of the slide making sure it comes in contact with all three smears. This step is tricky as it is easy to over- or under-decolorize. Do this for 10 seconds and IMMEDIATELY rinse, as in step 3, with a gentle stream of water.

6. Place the slide flat on the staining tray and dispense just enough Counterstain (0.6% safranin in 20% ethanol) to cover each smear. Let stand for 2 minutes; rinse with water as in step 3.

7. Blot dry using the bibulous paper package found in your orange drawer. Do not tear out the pages, just insert your slide and pat it dry.

8. Clean up your area; rinse your staining tray in the sink and leave it to drain upside down on paper towels.

9. Observe your stained microbes microscopically following the correct procedure for using the the oil immersion objective on your compound brightfield microscope. The directions for using the microscope are found in the protocol section of this wiki: BISC209: Microscopy

Use of the Compound Light Microscope

Activity: View your stained bacteria.
Refer to the directions for using your compound brightfield microscope BISC209/S11:_Microscopy found in the Protocols section. Today you will use only the 10x and 100x objectives. Remember also to read and follow the directions for care of this precision instrument (particularly on how to avoid getting immersion oil on any objective other than the 100x oil immersion lens). Be aware that there would be no field of microbiology if there weren't good, functioning microscopes to view this unseen world.

Gram Stain Results by use of Selective Media

Activity: Performing a Spot Inoculation Technique on Selective Media to Assess Gram Characteristics
Since you only have about 1/8 to 1/4 of the original colonies used in the activities above for each organism left, be judicious when you "confirm" the Gram reaction and check for contaminants by spot inoculation on solid selective and differential: eosin methylene blue (EMB) and phenylethyl alcohol (PEA) media. Consult Culture Media: Use of Selective & Differential media to confirm Gram stain. You should test all your isolates on both media and, if you do not see growth on either EMB or PEA, you should be able to explain this outcome as well as to explain the significance of growth on one and not the other medium.

Use a marker to divide the bottom of each plate into 4-8 sections and organize a labeling system in your lab notebook and on the plate so you can easily identify where you placed each of your soil isolates. You will spot inoculate the middle of each quadrant by taking a tiny amount of growth from the source isolate and inoculating with a single thin zig-zag line in the center of a section. See the illustration below of a plate testing 4 samples. If you do not have enough of the colony left to do this today, label the plates, and place them in your section of the cold room for use next lab when your stock subcultures have restored your supply of bacteria to test. Remember that bacteria reproduce asexually by binary fission so that if a colony comes from a single cell and you only use one colony or its descendants for all of your tests, you have used a genetically identical population (excluding spontaneous mutations) of cells for all of your tests over the semester.

Fig: 5C-1. Testing of multiple isolates in one plate can be accomplished by dividing a plate into 4 (OR MORE) sections. Be sure the inoculum is placed in the center of each section and that you check the plate for growth regularly.

Cultured Bacterial Isolate Characterization by Metabolic and Physical Tests

Testing for Antibiotic Production

Start the Testing for Antibiotic Production.

Many microbes secrete antimicrobial compounds to help them compete with other microorganisms for habitat. Some of the bacteria that are common antibiotic producers are the Actinomycetes (including Streptomycetes species), many of the Bacillus species, and the fruiting myxobacteria, to name just a few among many, many antibiotic producing bacteria. You can also test for the opposite: the sensitivity of your soil organisms (or known stock bacteria) to manufactured or secreted antibiotics.

(This testing will take 3 weeks.)
Week 1:
Identify how many potential antibiotic producers you might have. Definitely test any isolates that grew on glycerol yeast extract media are likely to be spore formers such as Actinomycetes, Steptomyces, or Bacillus. Test at least four isolates but there are extra plates if you would like to test more than 4. Do not test suspected Hyphomicrobia as they might not grow on the medium we will use today and you will use these later today in the Nitrogen test protocol. Since the soil is the main source of microbes that supply the world's antibiotics. It's possible that you might discover the next great antimicrobial drug and get very rich by selling the patent for your discovery to a drug company. Remember that the discovery of penicillin was completely accidental.

Using aseptic technique, transfer an isolated colony (possible Streptomyces, etc.) that's likely to be an antibiotic producer to a small clear glass tube containing 500μL of sterile water. Compare the turbidity to the #5 MacFarland standard tube that your instructor will provide. If it is too dilute, add more bacteria. If too concentrated, add more sterile water. Vortex to mix. Using a sterile swab, dip the swab in the diluted bacteria and make an inoculation (as shown below) down the middle of a plate of nutrient agar. Make a second plate exactly like the first for each isolate to be tested. Label them carefully and incubate the plates for ~1 week at RT.

Record all your results and observations.

CLEAN UP

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.

Assignment

Choose 8 of your team's isolates to test in the interaction assay to be started in Lab 7. Choose isolates that grow well on nutrient agar. 1-3days (depending on how fast your organisms grow) prior to Lab 7, please inoculate each of your team's chosen isolates into 2 tubes of sterile nutrient broth (16 cultures per team total). In addition, everyone should start a new isolation streak plate of EACH of your isolates on nutrient agar OR on the medium on which it grows (USE DMM for the Hyphomicrobia). In Lab 7 these cultures will be used for the quorum sensing assay and tests for nitrogen cycle role.

Write a summary of the theory behind the following techniques that we used to identify our bacterial species by molecular tools: genomic DNA isolation, polymerase chain amplification of part of the 16s rRNA genes, use of the Zero Blunt® TOPO® PCR Cloning Kit to create a library of unique plasmid vector with our 16S rRNA gene inserts and then select, One Shot® TOP10 Competent E. coli Cells that allowed us to select and separate our 16S rRNA genes for sequencing, and DNA sequencing by the newer fluorescent-labeled ddNPTs chain -termination (Sanger) method. Directions found at: Lab 6 Assignment: Assignment: Theory Summary

Links to Labs

Lab 1
Lab 2
Lab 3
Lab 4
Lab 5
Lab 6
Lab 7
Lab 8
Lab 9
Lab 10
Lab11
Lab 12