LAB 5: Soil Microbial Diversity &Function
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 a background reducer, 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. However, when one of our pcr products is ligated into the vector, the ccdB gene is disrupted, enabling these recombinant colonies to grow while other non-transformants do not. 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). Colonies that are transformed with "empty" vectors will not be selected out by plating the colonies on media with kanamycin since the kanamycin resistance gene will be expressed from the empty plasmid vector. 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. This means that cells containing a plasmid vector with our 16s RNA gene have this disruption of LacZ and ccdB gene regulation and will not be killed by absence of DNA gyrase. 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 only clone two pcr products/habitat (one per sampling site). Choose one pcr product per pair to use for cloning and transformation today. Choose the best initial genomic DNA concentration IF that pcr product had good amplification. If the amplification was unsuccessful or weak (judged by staining intensity on the gel), then use the most successful 16s rRNA gene amplification in each habitat sample.
Procedure: Add the reagents in this order!
1. Add 2 μl of PCR reaction 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.
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 to gather:
In addition to general microbiological supplies (e.g. petri dish with ethanol, glass spreader or sterile glass beads), you will need the following reagents and equipment.
• TOPO® Cloning reaction from Performing the TOPO® Cloning Reaction
• S.O.C. medium at room temp.(included with the kit)
• 42°C water bath
• warm Luria-Bertoni (LB) plates 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
• 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 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 and habitat, 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 neither of the transformations from your habitat 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.
Transformation Media Recipes
(may be stored at +4°C or
0.5% Yeast Extract
10 mM NaCl
2.5 mM KCl
10 mM MgCl2
10 mM MgSO4
20 mM glucose
0.5% yeast extract
1% NaCl <BR.
2% agar for solid medium, none for broth
50mg/L Xgal (optional)
Part III: Culture-Dependent Isolation & Characterization of Soil Bacteria: Obtaining Pure Cultures of Isolates
By this week we hope that you have pure cultures from your enrichment and isolation protocols. If you have isolated, pure colonies on your isolation streak plates, congratulations!. Make sure that these potentially pure cultures have colonies that look like the original source colony and that all of the colonies look the same (they can be of different size). If you are having trouble obtaining well isolated colonies, consult with your instructor. You may need to return to the original plates, to try isolation streaking again.
Your goal is that each person will isolate and identify 1 unique bacterial strain from each enrichment/isolation process. This means that each person will characterize 3 unique isolates. Coordinate with your partner and other classmates to try to isolate colonies that look different from each other. It would be great if everyone in the class identified different bacteria. That may not be happen, but it will make your papers and poster presentations at the end of this project more interesting if we have a wide variety of bacterial genera and/or species.
If you have pure cultures now, continue with the Activities that follow--if not, wait until you do have well isolated colonies in a pure culture. If you are unsure, consult with your instructor.
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.
Confirming your Gram Stain Results by use of Selective Media
Activity: Performing a Spot Inoculation Technique on Selective Media to Confirm Gram Stain Results
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.
Preparation for Next Lab
A battery of tests for metabolic and physicial characteristics of your isolates will continue next week. You will use the pure cultures that you have started this week or their descendants. Depending on the test, you will need a newly grown liquid broth culture, or an isolation streak plate culture. You will need to plan ahead to prepare the appropriate cultures so that they will be ready to use when needed. Try to use log phase cultures. The number of hours it take from inoculation until a bacterial culture moves from log to stationary phase depends on its generation time, the conc. of the inoculum, and other factors. If you have a reasonably fast growing culture, to inoculate your carbohydrate media with a log phase culture, you should make a subculture into an appropriate broth medium about 3-8 hours before you inoculate the test medium. Keep track of how fast each of your soil bacteria grow, on which media,and at what temperature. Since this is an investigative lab with no pre-designed outcome, success will require considerable planning and organization as well as copious notetaking.
Your most important resource for looking up information about your isolates will be the reference manuals: THE PROKARYOTES and Bergey's Manual. Wellesley College has these valuable reference books available in electronic form. Link to the electronic edition of | The Prokaryotesthrough Springer ebooks.
Link to the electronic edition of | Bergey's Manualsthrough Springer ebooks.
Use these reference resources to decipher the meaning of the results of the morphologic, metabolic, physical and other tests your perform. It is unlikely we will be able to provide ALL the tests you might wish to perform to identify and evaluate the various roles your soil bacteria may play in its ecosystem, but we will obtain interesting information. The DNA sequencing analysis of the 16s rRNA genes from these isolates should provide their identity, often to the genus and species level.
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 part of the Results section of your final paper on Bacterial Diversity in a Soil Community: Roles and Relationships
Please use the community physiological profiling data you have collected so far (including carbon source analysis and evaluation of nitrogen cycling, phosphate, cellulose, and starch processing bacteria in your soil community) to design and present effective figures with appropriate legends that show a new reader how your data answers one of the parts of our experimental question: how do the bacteria in a particular soil community function co-operatively?
Refer to the appropriate sections in the extensive handout, "Guidelines to Scientific Writing" found in the Resources section of the wiki. Your instructor is also available for guidance, and there are science writing peer-tutors, hired and supervised by the Writing Program, available for writing help. See the Writing Program web page for hours and availability or to schedule an appointment at | http://www.wellesley.edu/Writing/Program/tutors.html.
Continue monitoring and following the appropriate protocols to isolate and characterize a few of the culturable bacteria that make up your soil community. In Lab 6 and Lab 7, you will doing most of the assessment of your isolates' physical and metabolic characteristics through a battery of tests and special stains, a few of which require some preparatory work. Familiarize yourself with the tests and stains you will preform. Make sure you have outlined the protocols in your lab notebook and started any necessary cultures on appropriate medium so that they will be ready to use in lab at the appropriate time:
Links to Labs