January 21, 1998


Today we begin a four week unit designed to investigate the diversity and antibiotic resistance in bacteria (E. coli). Conceptually, the unit is divided into two sections:
1) A survey of the diversity of the gut bacterial ecosystem, and an investigation of the frequency of antibiotic resistance in a natural population of E.coli isolated from the lower G.I. of Yale undergrads, graduate students, faculty (and a few others).
2) Characterization of the plasmid diversity present in natural E. coli isolates, of the molecular details of plasmid-borne antibiotic resistance, and of the transmissibility of plasmids conferring antibiotic resistance.
*At the end of this document you will find a flow-chart summarizing the sections of this lab. Note that weeks three and four require you to come to lab outside of the normal scheduled class time.
This lab uses naturally occurring bacteria and weakened laboratory strains of E. coli. From the outset, we are selecting for E.coli isolates, and we are very unlikely to find anything else. However, some basic precautions need to be observed when working with these bacteria:
1) Wash your hands before and after you work in this lab.
2) Wear gloves unless we tell you otherwise.
3) Absolutely no eating or drinking of any sort in the lab. Get rid of the candy and chewing gum as well.
4) Absolutely no mouth-pipetting of any sort.
5) Do not discard anything -- swabs, tips, pipettes, plates, gloves -- in short, anything, into the regular trash. All trash goes into the specially designated containers, which will then be autoclaved before disposal.
6) Do not discard any liquids down the sink. Leave them in their containers (test-tubes, flasks, etc...) where they will then be autoclaved before disposal.
7) If you have (or recently had) any kind of intestinal infection, stomach flu or equivalent, let me or one of the T.A.'s know immediately.
8) Write all relevant information on the bottom of the plate, not on the cover. Covers get lost and shuffled around, and you can easily make a critical mistake. Label tubes and plates before you use them-- it is easier that way.
Today's lab begins our characterization of the bacterial isolates and provides an initial screen of the frequency of antibiotic resistance.

You will receive a handout explaining how to isolate single bacterial colonies, plate dilutions of bacteria and use a Pipetman microliter pipette. Some important things to remember:
* Do not adjust Pipetman out of its normal range (e.g., do not use a
P-20 to pipet 25 ul).
* Incubate and store plates inverted.
* Do not dig into agar when toothpicking colonies.
We will be using five different types of media in this unit. Each of them provides some specific information about the biochemical and metabolic behavior of the microorganisms we grow. In addition, these media may select for (or against) particular types of bacteria. Keep in mind that from the outset, we are selecting against anaerobic growth, a prevalent microenvironment in the human gut.
LB (Luria-Bertani): This is a rich medium that supports the growth of many species of Enterobacteriaceae. It does not select specifically for E. coli.
ML (Minimal Lactose): This medium uses small quantities of the sugar lactose as the main energy and carbon source. Many species of bacteria are not capable of transporting or utilizing lactose as a sugar source, and may also require additional components in the medium that they cannot synthesize on their own. E. coli is, for the most part, able to utilize lactose.
NaCi (Sodium Citrate): This medium contains citrate as a carbon source, which inhibits E. coli growth. If a colony grows on ML but does not grow on NaCi then it is most likely E. coli.
(Sheeps) Blood agar: This is a rich medium that allows the growth of many Enterobacteriaceae. It does not contain a colorimetric indicator which might interfere with the BBL Indole and Oxidase tests.

Collecting the field populations
We have given you collection swabs that are fairly self-explanatory, but here are a few pointers:
1) The best time to collect is in the morning, since your bacterial fauna has had all night to grow.
2) Don't be precious, and swab where you need to swab. The swab should no longer be white if you have done this right.
3) Replace the swab in the sleeve tightly, and break the ampule at the end of the sleeve by crushing it. The ampule contains a buffer that will keep your bacteria happy until you get to lab.
4) If you collect the sample the evening before, keep the swab refrigerated (if possible).
5) Label the sleeve with your name.
Estimating bacterial densities using serial dilutions
1) Bring your swab to lab.
2) Remove it from the sleeve and place it in a tube containing 1 ml. of dilution medium (DM). Vortex gently for 30 sec. in order to suspend the bacteria on the swab (make sure not to splash!). Remove the swab and discard it in red bio-waste bucket.
3) Prepare three dilutions of your original sample, using new tubes of DM. Make a 10-1 dilution (110 ul of the original in 1 ml of fresh DM), a 10-2 dilution (110 ul of the 10-1 dilution in 1 ml of fresh DM) and a 10-3 dilution (110 ul of the 10-2 dilution in 1 ml of fresh DM). Make sure to vortex, and be careful not to splatter.
4) Spread 100 ul of the 10-1, 10-2, 10-3 dilutions onto an ML (minimal lactose) plate, and make sure you label the plates carefully (name, date, dilution). ML is a growth medium that selects strongly for E. coli. The T.A.'s will show you how to do the spreading.
5) Spread 100 ul of the three dilutions onto an LB (Luria-Bertani) plate, and make sure you label the plates carefully (name, date, dilution). LB is a rich growth medium on which most everything will grow.
6) Spread 100 ul of the three dilutions onto Blood agar plates, and make sure you label the plates carefully (name, date, dilution).
7) Give all the plates 2-3 min to dry, bundle them together with your name clearly visible on the tape, and place them, INVERTED, in the 37oC incubator. Keep your DM tubes (labeled) in the 4oC refrigerator, just in case.
9) Next lab meeting, we will count the colonies that have grown on the LB and ML plates, calculate the density of bacteria in the sample, and begin the characterization of antibiotic resistance in your bacterial sample.

Wed. Jan 28th
Estimating the density of E.coli.
We will calculate a rough estimate of the original bacterial density by using the technique of serial dilution, where a standard aliquot of the initial DM is diluted to concentrations of10-1 ,10-2 and 10-3, poured out onto LB and Min Lac plates, spread, and allowed to grow overnight at 370C. Use the dilution that results in an easily countable number of individual colonies.
Original bacterial density is calculated by counting the number of colonies on a plate and multiplying by the dilution factor.
Making the master plate

Bacterial genetics depends on the use of genetically homogeneous single colonies, which are followed over time and characterized in various media. We will be preparing a master plate on ML, using a grid that permits single colonies to be grown in isolation. You will be toothpicking 16 spatially isolated colonies from the ML dilution plates onto this master plate (use a different toothpick each time) and incubating the plate overnight at 37 oC. Each individual colony will be assigned a number, and that number will remain constant throughout our experiments. Hang on to this plate carefully, you will be returning to it throughout this unit. Make sure that you can orient the plate in relation to the grid (make sure you know which colony "Colony 1" actually is).
Initial screening for antibiotic resistance: top agar and colony streaks.
We will be screening for resistance by making a bacterial lawn starting with a single colony. Subsequently, we will be placing small filter paper disks containing various antibiotics on that top agar lawn. We will then allow growth to proceed overnight, and check for zones of inhibition -- translucent zones surrounding the antibiotic disks where no bacterial growth has occurred. A zone of inhibition indicates that the bacteria are sensitive to that particular antibiotic. Conversely, if bacteria grow right up to the filter disk, the bacteria are resistant to the antibiotic on the disk (specifically to the dose of antibiotic on the disk) You will be preparing 16 bacterial lawns, each initiated with a single colony from your reference grid plate.
We will also ask you to prepare streaks of the individual numbered colonies for subsequent use. Streak two colonies on each Min Lac plate. Make sure that the streaks remain well isolated from one another, and are clearly labeled.

Once you have identified a potential colony from your ML dilution plates, you will be using it for three purposes:
1) to create an ML reference plate and a NaCi plate, containing the 16 chosen colonies, each growing in a grid square. The grid template will be handed out in class.
2) to create a confluent lawn with top agar on which we will place small antibiotic disks in order to assay resistance.
3) to streak out individual colonies onto ML plates for further characterization.
Here is what you do:

* take a sterile toothpick, break it in half (your hands are gloved, of course)

* touch the colony with the toothpick (scrape the colony, but do not puncture the agar)

* place an ML plate on the grid we will supply. Open it briefly, and touch your toothpick to the ML plate at the appropriate grid position (1st colony in position 1, 2nd colony in position 2, etc...). Make sure your plate is labeled so you can orient it on the grid. Here, again, you want to touch the toothpick to the ML plate, not pierce the agar.

* touch the same toothpick to the appropriate position on the NaCi plate.

* place the same toothpick into an Eppendorf tube containing 500ul of DM. Close, vortex briefly to suspend the cells remaining on the tip of the toothpick into the DM.

Making Top agar tubes:
*You will be pouring your own top agar tubes from a stock kept at about 65oC. Prepare 4 top agar tubes at a time (no more because they will solidify before you pour them) with 3 mL of top agar in each.

* take 200ul of the DM and add it to the top agar, after it has reached 50oC (you don't want it so hot that it kills the bacteria).

* swirl gently and pour out onto a phage plate. Distribure the top agar evenly over the surface of the plate by swirling. Cover plate.

* after the top agar solidifies, dispense the antibiotic disks onto the surface of the agar. Make sure the dispenser does not touch the agar! If the disks are not sitting properly on the surface, tamp them down with a sterile toothpick.

* sterilize a transfer loop, place the tip into the DM medium and use the streaking technique explained in the handout you received to streak ML plates. Streak two colonies per plate.

Incubate all plates overnight at 37 oC. The TAs will place the plates in the refrigerator on Thursday.

Antibiotic Disks

We are going to be testing for resistance to the following antibiotics: ampicillin, tetracycline, chloramphenicol, kanamycin, streptomycin, rifampicin, erythramycin, naladixic acid. There is a link to a web site which will explain how these antibiotics work, make sure you check this out.


WEEK THREE: Feb. 2nd, 3rd, 4th
This week you conduct the initial phenotypic survey of a sample from your E. coli population by scoring single colonies for their resistance or sensitivity to various antibiotics. We also begin to characterize the nature and molecular identity of that resistance. We will be asking the following questions:
1) Is the antibiotic resistance carried on the chromosome, or is it instead plasmid-borne?
2) Is there additional variation within each class of antibiotic resistance?
3) Do bacteria isolated from human G.I. tract carry multiple plasmids? Is there variation within a bacterial population for the number and type of plasmids?
4) Can we experimentally isolate and identify the plasmid carrying the antibiotic resistance?
The first issue we will address is the location of the genes conferring antibiotic resistance. The bacterial cell carries genetic information in two principal locations-- the chromosome, and on plasmids, small, circular extrachromosomal elements. Our experiment this week will capitalize on the ability of plasmids to transfer from one cell (the donor) to another (the recipient) with relative ease. While a variety of mechanisms (transposons, etc...) allow genetic elements to move from the chromosome, the frequency of such events is far lower than plasmid transfer.

Monday Feb 2nd:
1) Determine the antibiotic resistance profiles for your 16 plates. If the bacteria are sensitive to an antibiotic, there will be a zone of inhibition, a clear ring, surrounding the disk where the bacteria were unable to grow. You may only have one or two resistance profiles, e.g., 8 colonies tested were resistant to Amp and Tet and 8 were resistant to Amp and Ery.

2) Write your resistance profiles on the board. If your profile is already written on the board, and there are more than five cultures of that profile in the shaker, then you will only set up one culture from your own ML streak plates. If you have a new profile then start 3-4 overnight cultures of that profile so people can share in your good fortune.

For example, lets say I count my plates and I have three resistance profiles: 1) 10 plates were resistant to Amp, Ery and Ra 2) 3 plates were resistant to just Ery 3) 3 plates were resistant to Ery, Ra and Tet. I look at the board and see that someone has already written: Amp, Ery, Ra and there are ten cultures with this resistance profile in the shaker. This means that I will only start one additional overnight culture picking a colony from my ML streak plates with the resistance profile Amp, Ery, Ra. I look at the board again and find that the profile Ery is written on the board and there are 8 cultures in the shaker already; same thing, I start one additional culture. I look at the board and notice no one has written the profile Ery, Ra, Tet on the board yet so I write it on the board and start 3-4 cultures from my ML streaks with this resistance profile. I start several cultures of my unique profiles so that there will be enough for people to share on Tuesday when we start our mating cultures.

Note: if you have a rare profile, you can start multiple cultures from the same ML streak plate (using a different toothpick for each culture and picking a different colony from the same ML plate).

3) Pick the colonies from the ML plates you streaked during week two by toothpicking a single colony and dropping it into a tube of 2 ml liquid LB. Do this for each different resistance profile you have. These overnight cultures are the plasmid donor cells. Place them in the shaker bath at 37oC.
4) The TA's will prepare overnights of CSH50nalr cells. This is an E. coli strain that exhibits chromosomally encoded naladixic acid resistance but does not have any plasmid-encoded antibiotic resistance. These will act as the resistance recipient cells.

Tuesday Feb 3rd:
1) You will test for the transfer of Amp, Chlor, Ery, Tet, Strep, and Kan resistance (note that we do not have Rifampacin plates so cannot test for resistance transfer). You will set up six mating cultures, one for the transfer of each antibiotic resistance phenotype. In the event that no one is resistant to Chrolamphenicol, you will test for the transfer of a single antibiotic resistance from two different profiles (e.g., test for transfer of Ery resistance from an Ery, Ra profile and from an Ery, Amp, Tet profile).

2)Find a culture in the shaker confering the appropriate resistance, and take one tube of CSH50nalr cells. Place 100 ul of donor culture (resistant cells) and 100 ul of recipient culture (CSH50nalr) into new liquid LB tubes. Leave them overnight, in the incubator at 37oC, without shaking (why does the shaker need to be off?).
3) Pick 6 colonies ("different-looking" colonies if you have them) from your Blood agar plates and start your BBL Identification kits following the directions on the BBL Crystal handout. Don't forget to start with the Indole and Oxidase tests.

Wednesday Feb 4th:
1) Today you will plate the mated cells and test for plasmid transfer. You will need to plate one TLN+antibiotic plate for each antibiotic that you are testing for transmissibility. For example, assume that your donor clone showed Erythromycin (Eryr) and Streptomycin (Strr) resistance. Your recipient clone (CSH50nalr) was Nalr. If the plasmid transfer occurred, and both resistance genes are on the plasmid, the result should be a CSH50 cell that is Nalr, Eryr, and Strr. You will test for the presence of these cells by streaking on a TLNE (Tetrazolium Lactose/Nalidixic acid/Erythromycin) and a TLNS (Tetrazolium Lactose/Naladixic acid/Streptomycin) plate. Only recipient cells into which the plasmid has transferred will grow on both of these plates.
2) Control plates: You should also streak a positive and negative control plate. The positive control will show that there are still viable CSH50nalr cells even if they did not receive additional antibiotic resistance during the mating. Which type of plate should you use? The negative control should be a TLNX plate, where X is an antibiotic that the donor clone was NOT resistant to. Chloramphenicol resistance is uncommon so this makes a good negative controm.
Incubate all plates at 37oC overnight. The TA's will take the plates out on Thursday and put them in the refrigerator.
3) Score your BBL identification plates following the directions at the end of this handout.

WEEK FOUR: Feb. 10, Feb. 11
1) You do not need to come in.
Tuesday Feb. 10:
Today we will start growing cells for a plasmid mini-prep we will do tomorrow.
1) Score your TLN+antibiotic plates.

2) Start 8 overnight LB cultures:

a) Five cultures of successful donors; use donors with different resistance profiles if you can and start the cultures by picking a colony from the ML streak plates you made during week one.

b) Three cultures of successfully mated cells, i.e., antibiotic resistance was transferred. Pick two different matings if you can where different resistance was transferred. Start the cultures from the TLN+antibiotic plate. These cells should now contain the plasmid that confers resistance.

For all of the above, you can either use the loop (after flaming and cooling it) to pick off the colonies, or you can use a sterile toothpick which you touch to the colony and then drop into the LB (remember to use gloves). Label, label, label the LB tubes before you start, or all will be chaos.

Wednesday Feb 11:
Over the last two weeks we have constructed a fairly detailed phenotypic characterization of individual bacterial colonies. We now undertake a further description at the molecular level. This lab is devoted to the isolation of plasmids from the various E. coli isolates we have worked with thus far.
We will be using Quiagen vacuum minipreps to isolate plasmid DNA. In broad terms, we will first concentrate the cells grown overnight, resuspend them in buffer and lyse them (break them open) by the addition of detergent (SDS), causing them to release their DNA. We then separate this mixture of protein, lipids and DNA by precipitating the proteins and lipids and filtering them out. The supernatant is passed through a filter that binds DNA which is eluted from the filter with 50 ul of water.
You will receive a handout expaining how to use Quiagen spin mini-preps. Here are some additions to the protocol and tips on working with DNA:
1) Wear gloves at all times. Your carry powerful DNAses on your skin and in body fluids, and they can easily degrade your DNA preparations.
2) Since we are isolating large plasmids (some may be > 150Kb), you must be gentle. Once we lyse the cells, DO NOT VORTEX, or the plasmids will shear. Similarly, if you are drawing any DNA though a pipette tip, do it slowly and gently.
3) Take your time. Make sure that you know what you are doing, and what the next step is, before you start. Make sure that all the materials and solutions you need are at hand (this is a lot like cooking).
4) Keep your isolated DNA on ice, unless specified otherwise. The stray enzymes that can degrade your plasmid preps are all slowed considerably at 4oC. Keep in mind that we are using very small volumes, so it only takes a few seconds of holding the tube in your hand for the contents to be at 37oC.
5) If you are not sure, ask.
Concerning the Quiagen protocol:

* To concentrate your cells: place 1.5 ml of your overnight culture into an Eppendorf tube and centrifuge for 3 min at full speed (10,000 rpm). Be sure to balance the centrifuge by placing tubes of equal volume directly across from one another. After centrifuging, pour off the clear supernatant into a biological waste container and begin at step 1 of the Qiagen protocol.

* Do not wait more than a few seconds after adding buffer N3 to invert your tubes and mix the solutions. Waiting longer will allow some chromosomal DNA to remain in solution with the plasmid DNA.

Qiagen protocol:

*Before you begin label your Eppendorfs, TurboFilter and Qiaprep strips with your name and which sample is number 1, 2, etc. The TurboFilter (clear platstic) removes the proteins and chromosomal DNA from your preps and the Qiaprep column (blue plastic) binds plasmid DNA so it can be washed and eluted in a smaller volume.

1) Pipet 1.5 ml of your overnight cultures into 8 Eppendorf tubes and centrifuge 3 minutes at 10,000 rpm. Pour off liquid into red waste bucket.

2) Resuspend pelleted cells in 250 ul of buffer P1 by pipeting up and down. Close the tube and vortex for 30 seconds until there are no clumps.

3) Add 250 ul of buffer P2 to each sample and invert tubes gently 4-6 times. Incubate tubes at room temperature for 5 minutes.

4) Add 350 ul of buffer N3 to each sample and invert tubes gently and immediately 4-6 times.

5) Pipet the lysates from step 4 (850 ul per sample) into the wells of a TurboFilter strip. Each strip has 8 wells and you need to label the strip with your name and which sample is number 1, 2, etc. DO NOT ACCIDENTALLY PIPET YOUR SAMPLES INTO THE QIAPREP STRIP OR THE PREP WILL BE RUINED!

6) Place your Qiaprep strip in the vacuum manifold underneath your TurboFilter strip containing your lysates so that sample 1 in the filter lines up with sample 1 in the Qiaprep. When the manifold is full (6 students per run) a TA will turn on the vacuum and the lysate will be sucked through the filter into the Qiaprep.

7) Discard the TurboFilter and place the Qiaprep strip (which now contains your plasmid DNA) in the top section of the manifold.

8) Add 1 ml of (wash) buffer PB to each well and apply vacuum.

9) Add 1 ml of buffer PE to each well and after PE flows through apply vacuum for an additional 5 minutes to dry membrane. This is important, if PE is left in your filter when you elute your DNA then your DNA sample will float out of the wells in your agarose gel = big problem.

10) Place blue 1.2 ml collection tubes underneath your Qiaprep 8 strips in the manifold (make sure they are labeled 1-8, etc.). Add 50 ul of elution buffer EB to the center of the Qiaprep membrane (make sure it is absorbed into the membrane and does not stick to the side of the tube).

11) Apply vacuum for 3 min to collect DNA. After collection, transfer your DNA samples to labeled Eppendorfs and place on ice. The DNA is now ready to run out on a gel.

Running plasmid profiles on agarose gels.
The protocol you followed should have yielded a solution containing all of the plasmids carried in the E. coli colony you grew up overnight. Ideally, these plasmids are still intact -- they are closed, double stranded circles of DNA that may range in size from 3 to 150 Kb. We are going to analyze these plasmids using the technique of gel electrophoresis. This technique involves placing the plasmid DNA onto an agarose gel matrix; the gel is then subjected to an electric field. The DNA is negatively charged and will therefore move through the gel towards the + pole. The rate at which the DNA migrates depends in part on the size of the molecule (other factors include the conformation of the DNA molecule and the base composition of the DNA molecule). The concentration of the medium (agarose or acrylamide) through which the DNA is moving also plays an important role.
In this case, we will be resolving the plasmids using a 0.7% agarose gel, prepared in TBE (Tris, Borate and EDTA) buffer. This gel contains highly toxic Ethidium bromide which allows the DNA to be visualized under UV light: wear gloves at all times when handling it. The gels are also very fragile so be careful unwrapping them.
Place the gel in the running chamber (containing 1X TBE buffer). If there are extra wells, practice loading wells by using the blue loading dye we have prepared for you. Get a feel for the process before you load your actual plasmid DNA.

Once you feel comfortable, mix in the appropriate amount of loading dye (4 ul of loading dye for 25 ul of plasmid prep DNA) and load onto the gel. When the T.A.'s give you the go-ahead, load the plasmids onto the gel. BE SURE TO RUN THE DONOR AND RECIPIENT PLASMID PREPS IN ADJACENT LANES. This will make it easier to compare the plasmids that were transferred. The TA's will let the gel run the appropriate amount of time, take a picture, and post it on the EEB Homepage under Student data.
1) Ethidium bromide is highly toxic. Wear gloves.
2) You are still working in a bacterial lab. Wash your hands.
3) These agarose gels require substantial current to run. Be very sure that the current is off before you load DNA or touch the gel or the buffer.
4) The only way we can figure out the size of the plasmids is by measuring their mobility relative to known size standards. Make sure that every gel has at least one lane devoted to size markers, which the T.A.'s will provide

BBL CrystalTM Identification Kits:
You will be using these kits to identify five bacterial colonies from the Blood agar plates you made during Week One. Each colony will be tested for i) the presence of tryptophanase (indole reagent test), ii) the presence of indophenol oxidase (oxidase test) and iii) the colony's abilitiy to utilize various nutritive substrates. The results of the three tests are entered into a computer program which determines the bacteria species.
Pick big colonies to identify because each colony will be used to streak one filter paper for the Indole test, one filter paper for the Oxidase test and also used for the substrate utilization test. You will receive a handout of instructions for the substrate utilization test and here are the instructions for the Indole and Oxidase tests:
BBL DMACA (Indole reagent) Instructions:
1) Squeeze dropper in the middle to break ampule inside and tap on tabletop to mix reagents.
2) Moisten a piece of filter paper with a few drops of indole reagent.
3) Remove a well-isolated colony from your culture plate with a toothpick and smear on moistened filter paper.
4) Observe for a blue to blue-green color within 2 min for a positive reaction for indole production. A negative result displays no color change or a pinkish tinge.
BBL Oxidase Instructions:
1) Squeeze dropper in the middle to break ampule inside and tap on tabletop to mix reagents.
2) Moisten a piece of filter paper with a few drops of indole reagent.
3) Remove a well-isolated colony from your culture plate with a toothpick and smear on moistened filter paper. Do not add excess reagent, it may cause the reaction to fade on oxidase-positive organisms.
4) Positive reactions turn the bacteria violet to purple immediately or within 10-30 seconds. Delayed reactions should be ignored.