CE 580: Biomolecular Tools for Engineers Lab Manual

Overview: Teams have been assigned with an effort to choose partners with complementary skills/background. Each team should choose an engineering relevant sample that they will focus on for the semester. During the course of the semester you will qualitatively and quantitatively describe the bacterial composition of this sample. General Guidelines in the Lab: Proper attire: Always wear gloves in the lab! This is both for your protection, and also to protect the samples from contamination from your skin. Remember, DNA is everywhere, you want to make sure you are studying the DNA from your sample, and not random environmental DNA. A lab coat or other lab smock is recommended to protect your cloths from spills. You may keep this in the lab for use during lab classes. Also remember to wear closed-toed shoes and long pants or skirts which protect your legs. Waste Disposal: Most waste generated in this lab can be disposed of in the regular waste bin. However, there are some exceptions noted below. These should be disposed of in the appropriate marked container for hazardous waste collection. Hazardous Chemicals: Formamide: This is used in small quantities in PCR and also in the making of the DGGE gel in order to lower the melting temperature of DNA. Acrylamide: This is the main ingredient in the DGGE gel, and is a known neurotoxin. We will only use liquid acrylamide, which eliminates hazards associated with inhaling the powder form. Using the pipeters: Pipeters are expensive tools that must be cared for properly. Each group has 4 pipeters with 4 volumes: 0.5-10 µl, 1-20 µl, 10-100 µl (or 200 µl), and 100-1000 µl. Always choose the appropriate pipeter for the appropriate volume. Set the volume on the pipeter and then take a tip with the pipeter. When taking sample, release the button slowly to draw sample into the tip. If you release the button too quickly, the volume will not be accurate, and may splash inside the pipeter. Always hold pipeter vertically, turning the pipeter sideways with sample in the tip will contaminate the pipeter and your sample. Finally, after taking the sample, check to make sure there are no bubbles in the tip before dispensing the sample. Dispense of the tip and use a fresh tip for each sample.

Team Assignments: Group 1 Rachel Hanson & Loona Borgohain Group 2 Sage Hiibel & Mary Jean Jones Group 3 Heather Storteboom & Catherine Hong Group 4 Ashish Sharma & Christian Lee Group 5 Maria Raynal & Rachael Kurkowski Group 6 Susan Yonemura & Arlin Ward Lab I: DNA Extraction and Quantification Objective: You will extract DNA from the sample of your choice using the QBiogene FastDNA SpinKit for Soil. This method employs physical disruption of the cells (bead-beating) followed by physical binding the DNA to a silica matrix, washing with an ethanol based solution, and eluting in purified water. Procedure: Selection of Sample: Each team should select a sample for DNA extraction. Examples include: bacterial cultures (relatively easy to extract), soil (relatively more difficult), sediment, bioreactors, or sludge. You may also choose a sample relevant to your graduate research. Make sure this sample is of interest to you, you will be studying this sample the rest of the semester. Bring with you enough sample to perform the extraction in duplicate. For soil or other solid sample, 0.5 g of sample is required per extraction. For culture, 1 ml is typically sufficient, but you will need to centrifuge it first, pour off the supernatant, and transfer the pellet to the extraction tube (For dilute cultures, up to 50 ml may be centrifuged). If you are unsure, ask the instructor or TA for advice on what sample and how much to bring. DNA Extraction: A. Disrupting the Cells:

1.) Do a duplicate extraction of your sample. Weigh the empty matrix tubes and record the weight. Add your sample to the “matrix tubes,” and record the volume with sample. Use a sterile instrument (such as a spatula or pipette tip) to transfer the sample into the tubes.

2.) Add 978 µL of phosphate buffer and 122 µL of MT Buffer to the matrix tube using the 100-1000 µL pipette.

3.) Place cap on tightly and secure tubes in bead-beater. Make sure that samples are balanced, as you would in a centrifuge. Place setting on “Homogenize”, and run for 3 minutes. Note: because of the force exerted in the bead-beating process, it is recommended that you place the bead-beater on the floor during sample processing.

4.) After bead-beating, centrifuge sample for 1 minute on highest setting (making sure samples are balanced).

B. Binding the DNA: 5.) Transfer the supernatant (the liquid forming the top layer) to a fresh 1.5 mL centrifuge

tube. Add 250 µL of PPS (protein precipitating solution). Mix by inverting the tubes 10 times, then centrifuge on the highest setting for 5 minutes.

6.) While centrifuging, label one 15 mL centrifuge tube (with blue cap) and add 1 mL of Binding Matrix (make sure to shake the binding matrix prior to adding). After centrifuging, transfer the supernatant to a 15 mL centrifuge tube, taking care not to disturb the pellet. Swirl the sample gently to mix. Do this several times to maintain the sample in suspension for 2 minutes.

7.) Set the 15 mL tubes aside and allow the binding matrix to settle. Once settled, remove 0.5 mL of the supernatant with the pipette tip and discard, taking care not to remove any settled binding matrix.

8.) Resuspend the sample by swirling and transfer 600 µL into a tube with a spin filter (make sure to label these tubes appropriately). Centrifuge the tube with the spin filter for 1 minute. The DNA should stay bound to the matrix, and the remaining solution will come down into the bottom of the tube. After centrifuging, open the tube, take the spin filter out with one hand, and with the other, discard the flow-through in the bottom of the tube. Replace the spin filter and add 600 more microliters of the suspended binding matrix. Centrifuge, discard flow-through, and repeat with remaining sample.

C. Washing the DNA: 9.) Add 500 µL of SEWS-M to the spin filter. Centrifuge and discard flow through. After

discarding flow-through, centrifuge empty tube for 2 minutes to “dry” the spin filter. 10.) Transfer the spin filter containing the washed binding matrix to a fresh catch tube. Allow

to sit for an additional 5 minutes with the lid open to further dry the binding matrix. D. Eluting the DNA:

11.) Add 50 µL of DES (ultra pure water, DNA and pyrogen-free) and stir the binding matrix gently with the pipette tip. Be careful not to put a hole in the spin filter while stirring.

12.) Let incubate 2 minutes and then centrifuge for 1 minute on high. Check to make sure that the flow-through is clean, and none of the binding matrix passed through the filter (If this happens, transfer the sample to a fresh tube, being careful not to transfer any of the binding matrix.

13.) DNA is now extracted! Transfer each extraction to a freezer tube. Be sure to label this final tube appropriately for storage (Name, date, sample identification). Store sample at –20 ºC.

Lab 2: Electrophoresis and DNA Quantification Objective: You will verify the DNA extraction by agarose gel electrophoresis. This method will be used throughout the semester to visualize DNA. You will also quantify the extracted DNA by spectrophotometry and prepare dilutions of your DNA for subsequent PCR reactions. Agarose Gel Preparation:

1.) Set up the casting tray in the gel caster. Place the comb with the appropriate number of wells in place at one extreme end of the gel tray.

2.) Pour a 1.2 % solution of agarose in 1X TAE buffer (already prepared and melted at 55ºC) into the gel tray. Fill until about ¼ cm from the top of the space in the comb. If any bubbles form while pouring the gel you may pop them or push them to the side using a pipette tip.

3.) Set aside and allow the gel to solidify.

Loading and Running an Agarose gel: A. Setting up the Electrophoresis Unit

1.) Carefully remove the comb from the solidified gel by pulling the comb slowly upwards. 2.) Prepare 1 L of 1X TAE buffer solution by diluting the 50X concentrated TAE buffer with

DI water (eg, 2 ml of 50X TAE per 100 ml solution = 1 X TAE). 3.) Turn the gel tray containing the solidified gel so that the wells are closest to the negative

(black) electrode. [DNA has a negative charge, so will travel towards the positive (red) electrode once potential is applied.]

4.) Pour the 1X TAE solution over the gel so that it fills the wells in the gel, and also fills the electrophoresis chamber flush with the top of the gel.

B. Preparing the Samples and Loading the Gel You are going to load a total of four samples: the two samples that you extracted and two molecular weight standards.

5.) In a 0.5 mL tube mix 6 µL of blue loading dye with 2.4 µL of a 1:100 solution of SYBR Green in methyl sulfoxide (already prepared). SYBR Green is a DNA stain. It binds to double stranded DNA and fluoresces under UV light

6.) Now cut a piece of parafilm (about 2 inch by 4 inch should be sufficient) and place on the lab bench, parafilm side up.

7.) Using the 0.5-10 µL pipette, place four 1.4 µL “dots” of blue-green loading dye/SYBR Green solution on the parafilm (You should have some extra solution left because in step 5 you prepared enough solution for 6 dots to take into account pipetting errors).

8.) Now take 3 µL of the first molecular weight marker, and add to the first blue-green dot. Mix by pipetting up and down. Once mixed, take the sample and dye mixture back up in the tip.

9.) Load this sample into the first well. Do this by placing the tip gently about midway into the well, and slowly releasing the mixture into the well (Careful! Do not pierce the bottom of the well!). The blue loading dye contains glycerol, which makes the sample sink to the bottom of the well, and helps prevent the samples from coming out of the wells and cross-contaminating other wells.

10.) Repeat this procedure with the two DNA extracts, and finally with the second molecular weight marker.

C. Running and Visualizing the Gel: 11.) Once the gel is loaded, place the cover on the electrophoresis unit (red to red, black to

black) and plug in the leads to the power supply (red to red, black to black). Set the voltage on 90 v and press “run.” If you are running two gels from the same power supply, assuming that the resistance of each gel is the same, set the voltage on 180 v. Run until you see the blue dye move about halfway down the gel. At this time, turn off the voltage to the gel.

12.) In order to “see” the DNA, you need to look at it under UV light. Take the gel over to the imaging system. This system consists on a box with a camera located on top of a UV table. Turn on the UV light and take a digital image of the gel (the TA will explain how to do this during the lab session). If your extraction is successful, you should see something similar to Fig. 1. On your gel you will have two molecular weight standards on either side of your samples.

Fig. 1: Example of DNA extract run on agarose gel and visualized under UV light. 1.) molecular weight marker or “ladder” with top band of DNA = 2000 bp, followed by 1200 bp, 800 bp, 400 bp, and 200 bp. 2-6) Extracts of microbial community genomic DNA that is largely intact (no smearing). Quantifying the DNA: The brightness of the band of the extracted DNA on the agarose gel should give you an indication of the yield of your extracted DNA (brighter = higher yield). The 2000 bp band of the standard (the highest molecular weight) has a known mass of DNA per microliter (50 ng/µl),

1 2 3 4 5 6

which can be used to estimate the concentration of your DNA in ng/ µl. The integrity of the DNA can also be estimated (single band versus “smear”). Now we will use spectrophotometry to better quantify this yield. You may want to get this part started while you are running the agarose gel.

1.) Warm the lamp on the spectrophotometer for 30 minutes prior to use. 2.) Make a 1:100 dilution of your DNA extract, with a final volume of 500 µL (eg. 5 µL of

DNA extract added to 495 µL of D.I. water). You can do this in a microcentrifuge tube. You may use regular D.I. water to do the dilution (it is not necessary to use the ultra pure water, since you will throw away this sample after measuring it).

3.) Once the lamp is warmed-up, fill the quartz cuvette with the same water you used for the dilution. Zero the instrument. Note: Handle the quartz cuvette carefully, it is expensive! Only wipe the surface of the cuvette with kimwipes, other materials may cause scratches.

4.) Once the instrument is zeroed, check the samples. Rinse out the cuvette with D.I. water, “tap” dry on a kimwipe, and transfer the sample to the cuvette using the pipette.

5.) You will determine the absorbance at two wavelengths: 260 nm (DNA) and 280 nm (protein). A high ratio of 260/280 indicates that the sample is relatively pure with respect to protein contamination.

6.) Use the following formula to determine the concentration of DNA in your sample:

To calculate the concentration of genomic DNA in the dilution: A260 * 50 ng / µL = x ng / µL

To calculate the concentration of genomic DNA in the extract before dilution: x ng / µL * (500 µL / 5 µL) = y ng / µL

To calculate the total mass of genomic DNA in the extract: y ng / µL * 50 µl = z ng

Preparing dilutions of your DNA for PCR:

You will be using your DNA extract as a template for polymerase chain reaction (PCR) in subsequent laboratories. PCR is highly susceptible to the presence of contaminants in the DNA extract, which can inhibit the reaction. Typically DNA extract is diluted prior to PCR in order to dilute out inhibitors. Based on the concentration and purity of your DNA extract, you will need to do a dilution of your DNA before using for PCR. If the ratio of 260:280 is 1.5 or greater, then you have relatively pure DNA and you will not have to do a high dilution (1:3 and 1:5 are probably a good dilutions to try in this case). If this ratio is less than 1.5, then try a higher dilution (1:7, 1:10 or 1:20 for high concentration DNA). Do not dilute the whole DNA extract, instead- add the required volume of dilution water to a clean microtube and add the appropriate amount of DNA extract to this tube (typically 1-2 µl). For example, for a 1:5 dilution, add 4 µl of water to a microtube, and 1 µl of DNA extract. You should prepare two dilutions of each of your DNA extracts and store them at -20ºC until used for PCR next week. If you are unsure, show your gel and spectrophotometry results to the TA or professor to obtain guidance on what range of dilutions you should prepare.

Lab 3: Polymerase Chain Reaction (PCR) Objective: This week we will be doing PCR. This is a process by which targeted portions of bacterial genes may be selectively amplified to create millions of copies. This provides sufficient material for downstream analysis. We will be discussing PCR in a short lecture today and in Monday’s lecture next week. Today in lab you will use PCR to amplify a 200 bp portion of the 16S rDNA genes from the bacterial community DNA that you have extracted. This PCR reaction will target a ~200 bp hyper-variable portion of the 16S rDNA genes that is ideal for separation by denaturing gradient gel electrophoresis (DGGE- next week’s lab). You will use primers I-341F and I-533R to target this region (Fig. 1). The forward primer also contains a GC clamp that will help aid separation by DGGE in next week’s lab.

Fig. 1: Approximate position of PCR primers for PCR of 16S gene. Preparation: PCR is an exponential reaction. Therefore, it is highly susceptible to contamination by foreign DNA. Clean the surface of your working area with ethanol, and then with “DNA away” to sterilize the area and minimize any foreign DNA. Also clean the pipetters you will use with DNA away. As always, wear gloves. Each group has a Styrofoam container filled with ice. The PCR reagents should all be kept on ice as much as possible during reaction preparation. This reduces the activity of the Taq DNA polymerase during preparation. If the reaction mixture is not kept cold during setup, then this increases the likelihood of the formation of non-specific PCR products. Each group also has a set of “aerosol barrier” pipet tips. You will notice that these tips contain a white “plug” that helps minimize cross-contamination of aerosolized DNA between samples. Use these tips while setting up the PCR reactions. Setting up the PCR reaction with primers I-341F and I-533R: PCR reagents for each group have been aliquotted and stored in the freezer. Remove the reagents and allow them to thaw on ice. Table 1 summarizes the reagents and quantities per reaction needed. In the column on the far right, you should calculate and record the amount you need to add to your master mix for the number of reactions that you have:

16S rRNA gene ~ 1500 bp

341F 533R

5’

5’

3’

3’

Table 1: Summary of components for PCR with Primers I-341F and I-533R. Reagent Amount needed per

25 µµµµl reaction “Master Mix”: Amount

needed for N+1 Reactions (calculate)

Purified Water 13.15 µl 10X buffer (w/ 50 mM Mg++) 2.5 µl 5X buffer 5 µl dNTP (10 mM) 1 µl Primer I-341F (5 µM) 1 µl Primer I-533R (5 µM) 1 µl Taq Polymerase 0.35 µl

Totals 24 µµµµl The Master Mix contains all of the reagents needed for the number of PCRs you will carry out, 24 µl of which will then be aliquotted into PCR microtubes and finally one microliter of your extracted DNA (testing at least two dilutions of each of your two extracts prepared the previous week) will be added for a 25 µl total reaction. You will be carrying out 4 PCR reactions: 2 dilutions for one of the extracted DNA samples and one dilution for the second DNA extract as well as 1 negative control. You need to prepare extra Master Mix in order to account for losses and pipetting error. Therefore, prepare enough Master Mix for N+1 (5) reactions. After you have finished the calculations, and the PCR reagents have thawed, you may begin preparing the Master Mix. First, vortex all of the reagents in order to eliminate any concentration gradients which may have formed during freezing/thawing of the reagents. Then, add the reagents to a microcentrifuge tube in the order they appear on the table. It is a good habit to add in this order (least costly to most costly) so that if any mistake is made you do not have to throw away expensive Taq polymerase (one 200 µl tube = $350.00!). After you have added all of the reagents- vortex and place on ice. Now label the microtubes that you will use for the PCR reactions. Aliquot 24 µl of Master Mix into each of the PCR tubes. Finally, add 1 µl of the diluted DNA extract to each of the two sample tubes. To the negative control, add 1 µl of purified water. When all of the groups are ready- place the tubes in the thermal cycler. Until then, maintain the microtubes on ice. All of the samples will be run on the thermal cycler together- the program takes about 1.5 hours. Table 2: Thermocycler program for Primers I-341F and I533R: 94 ºC 2 minutes Initial denaturing step 94 ºC 15 seconds denaturing 52 ºC (reduced by 1ºC every two cycles until reaches 47ºC)

15 seconds Primer annealing

72 ºC 20 seconds extension Repeat step 2-4 for 35 total cycles

Amplification

72 ºC 7 minute Final extension step 4 ºC � Hold Making the agarose gel: While you are waiting for the first PCR reaction to finish you will make an agarose gel. The 1.2% agarose should already have been melted for you by the TA. Set up the gel casting tray for the appropriate number of samples and pour the mixture. Be ready to pop any bubbles that form with a pipet tip or needle. Load and Run the Agarose Gel: Once the gel has cooled and your PCR reaction is done, remove the comb. Prepare 1X TAE buffer from the 50X concentrated solution using DI water (you will need about 250 ml per gel). Pour the TAE buffer over the gel, filling the wells and the chamber until the buffer level is about even with the top of the gel. Load 3 µl of the PCR product onto the agarose gel, using 1.4 µl of blue loading dye with 1:100 SybrGreen and parafilm as you did for Lab 2. Don’t forget to load the DNA molecular weight standard (“ladder”). Load 2 µl of the standard. Run the gel at 150 V until the blue dye is about 1/3 to ½ way down the gel. Look at the gel under UV light to verify PCR product. The expected size of the PCR product is about 200 bp. If your reaction is successful, you should see a band at ~200 bp (see appendix for molecular weights of bands in size standard). Clean up: Save your PCR product(s) (well-labeled!) in the refrigerator, you will need this next week when we do DGGE. Also remember to replace your DNA extract and any unused PCR reagents in the freezer. Empty the buffer from the electrophoresis chambers, rinse with DI and place on the rack to dry. Appendix: Primer Sequences: 1I-341F (with GC clamp): CGCCCGCCGCGCGCGGCGGGCGGGGCGGGGGCACGGGGGGCCTACGGGIGGCIGCA 2I-533R: 5’-TIACCGIIICTICTGGCAC-3’

1 This primer contains a GC clamp at the forward end to help with resolution when running DGGE 2 I indicates inosine base- helps increase sensitivity of primer

Low DNA Mass Ladder Molecular Weight Standard:

Lab 4: Denaturing Gradient Gel Electrophoresis (DGGE)

Objective: You will resolve your ~200 bp PCR products (with GC clamp) on a DGGE gel in order to get a profile of the microbial community present in your sample.

Overview:

We will meet at Yates 314 for this lab. At least one group member needs to meet the TA on Tuesday to load the gel- we will figure out a time that works in class. Because we only have one DGGE unit, this lab will be a more of a demonstration than previous labs. Rather than working separately in groups, the six teams will run their samples together on two gels. Each gel can hold 14 samples, and each of the six groups has 2 to 3 PCR products (depending on how many worked). Also, because about 22 hours are required to run the gel, the TA will prepare the gels in advance and we will meet the day before that lab to load the samples (this part shouldn’t take too long). On Wednesday we will take a picture of the gels, cut the bands and then, we will prepare a gel together so that you can see how it is made.

Procedure: TUESDAY

Running the Gel:

Washing the Wells:

When we meet on Tuesday, the TA will already have prepared the gels and they will be in the electrophoresis chamber with 0.5X TAE buffer solution and heated to 57º C. This high temperature helps to denature the DNA. First you will rinse the wells in order to flush out any residual unsolidified gel which may be present in the wells. Washing the wells is a critical step for preventing the samples from degrading inside the wells. Wash the wells by filling a 10 mL syringe with the heated buffer and flushing each well individually. Flush each well each well 3 times.

Loading the samples: Add 10 microliters of blue loading dye directly to the PCR tubes with the 200 bp product and

mix by pipetting up and down. Before loading, the TA will turn off t the pump and rinse the wells one final time.

The TA will load the first sample, and then each group will load their samples in order. You

will load the gel in the order of your group number in the following order: 1.) negative control 2.) first sample 3.) second sample 4.) third sample (if available). You will use extra small-bore pipet tips that allow you to reach the bottom of the well with the sample. Make sure when you take the sample into the pipette tip that there is a bubble of air beneath the sample (by setting the pipetter volume higher than the actual sample volume)- otherwise capillary action will pull your sample out of the tip before you are able to position it in the well

Running the gel: After all of the groups have loaded their samples, the TA will run the gel overnight at 45

volts. We will wait to turn the pump on until the samples have visibly migrated into the gel. The pump recirculates the buffer in the top chamber of the unit so that it maintains contact with the electrodes. Turning on the pump too early, however, may disturb the freshly loaded samples. We will run the gel for 21 hours and 10 minutes at 45 volts.

WEDNESDAY

Preparing the Gel Stain:

Instead of SybrGreen we will use SybrGold nucleic acid stain. This stain is much more sensitive than SybrGreen or ethidium bromide and has the advantage that it does not emit background fluorescence when not bound to the DNA. We will prepare a 1:10,000 dilution for each gel (20 uL in 200 mL) in 1X TAE buffer. We will keep these solutions away from light until ready for use (SybrGold is photosensitive).

Staining the Gels:

Once the gels are finished, we will remove them from the unit, cut off the wells, notch the top right corner (for orientation), and place them in two trays that contain the buffer with the gel stain. We will then place the trays on an orbital shaker (covered) for 15 minutes. While the gels are staining, we will get the imager ready. Also, each group should label five 1.5 ml microcentrifuge tubes. We will put the gel slices in these tubes when we cut the gel.

Preparing a DGGE Gel:

Important note: Before taking a picture of the DGGE gels that we loaded on Tuesday we are going to start preparing a DGGE gel so you can see how this is done. The preparation of a DGGE gel requires some waiting time to let the gel solidify. We are going to use the waiting time to take the picture of the DGGE we prepared on Tuesday and cut the DGGE bands.

Setting up the Glass Plates: First the TA will demonstrate how to prepare the glass plates for pouring the gel: First they

are coated with SigmaCote or RainX using a Kimwipe (Warning: SigmaCote is volatile and it is dangerous to inhale the fumes- we will do this in the fume hood). Then the spacers are put in place and the plates are clamped so that the spacers and the bottom of the glass plates are perfectly flush. After this the plates will be secured in place and the gel will be prepared.

Mixing the gel solutions: We will keep the gel solutions on ice so that it does not solidify until we are ready. One 50

mL centrifuge tubes will be labeled “H” (for high-density solution) and another will be labeled “L” (low density solution). The following will be added to the tubes:

30% to 50% Denaturing Gradient

H L 100% denaturing solution 9 mL 5.4 mL

0% denaturing solution 9 mL 12.6 mL Blue dye 320 µL ---------

The 100% denaturing solution contains 7M urea and 40% vol./vol. formamide in 1X TAE

buffer and 8% acrylamide/bis solution (the main component of the gel). The 0% solution contains only acrylamide/bis in 1X TAE. The blue dye serves as a marker to distinguish the tubes and also so that the gradient can be visualized when it is poured. Danger! Acrylamide is a neurotoxin, and should not be allowed to come into contact with the skin!

Now we will prepare a 100 mg/mL solution of Ammonium persulfate (APS) in DI water.

This will be added later to help solidify the gel. Once this is made, the syringes and the gradient pourer will be put in place. It is important to have everything in order before pouring the gel- once the solidifying agents are added- you only have about 10 minutes to pour the gel before it begins solidifying.

Once we are ready to pour the gel, we will add about 2.5 mL (can be approx) of 100%

denaturing solution to a separate centrifuge tube. This will be used to form a seal at the bottom of the gel. Add 7.5 microliters of the APS solution and 5 microliters of TEMED to this tube- vortex- and immediately fill a syringe and dispense enough of the gel solution between the plates to form a thin layer at the bottom of the glass plates. This must be done very quickly or the solution will gel in the syringe. Rinse the syringe out when done.

Now add 18 microliters of APS and 18 microliters of TEMED to each of the tubes labeled

“H” and “L”. Fill each syringe with the appropriate gel solution and position them on the gradient pourer (blue tube on the right side). Once in position, the syringes will be connected by a three-way connector and on the third end an 18 gauge needle will be connected. Finally, the

cam of the gradient pourer will be turned slowly in order to avoid the formation of any bubbles. You will notice that the blue solution pours faster at the beginning than the clear solution- this is how the gradient is formed. Once the gel solution reaches the top of the plates- a 1 inch spacer will be put in place and the gel will be placed in a warm place to aid solidification. Now, we need to let the gel solidify for 1 to 1.5 hours. In this time we are going to take the pictures of the DGGE gels and cut DGGE bands.

Imaging the DGGE Gel: We will carefully transfer the first gel from the stain solution to the imager. The gel is very

fragile and can easily tear if mishandled. Once on the imager- we will take a digital image of the gel. After this- we will transfer the gel onto the cutting tray in order to avoid damaging the surface of the UV table with the razor blades. Each group will then take turns cutting dominant bands from the gel. The key to cutting the bands is to avoid the edges and only cut the central 1 mm square portion of the band. Pick at least 5 bands and transfer to the microcentrifuge tubes. Add 36 microliters of sterile water and make sure that the gel piece is pushed all the way to the bottom of the tube and submerged in the water. Then, we will do the same with the second gel. Caution! Make sure to wear the UV shield and protect all exposed skin from the UV light! Trust me- it is possible to get a painful burn from the UV light…

We will store the gel slices in the freezer until we are ready to re-amplify them for

sequencing.

Preparing a DGGE Gel-continued: Once the DGGE gel solidifies- we will prepare the “stacking gel” or the layer of the gel with

the wells. This is made just as the bottom of the gel was made- only with 0% denaturing solution rather than the 100%. First, we will remove the spacer and pour off any residual unsolidified gel. After this, we will prepare 5 ml of the stacking gel solution (with 10 microliters of TEMED and 15 microliters of APS). After vortexing and quickly adding the stacking gel solution to the top of the gel with a syringe- the 16 well comb will be put in place and allowed to solidify for about an hour.

PM

Fig. 1: Image of DGGE gel comparing microbial communities present in acid mine drainage remediating communities.

Lab 5: Polymerase Chain Reaction (PCR)- Preparing for cloning and sequencing of DGGE bands Objective: You will use PCR to amplify the 16S rDNA genes of the bacterial community DNA. You will use this PCR product for cloning. While this PCR is running, you will set up a second PCR using the DGGE bands that you cut last week as a template. The figure below indicates the position of the primers (8F and 1492R) that will be used for amplifying the full-length of the 16S rDNA genes in your sample, with respect to the primers you used for the DGGE PCR in Lab 3.

Preparation: Remember to clean the surface of your working area with ethanol, and then with “DNA away” just as you did in lab # 3. Also clean the pipetters you will use with DNA away. As always, wear gloves. Each group has a styrofoam container filled with ice. The PCR reagents should all be kept on ice as much as possible during reaction preparation to reduce the activity of the Taq DNA polymerase during preparation. Use “aerosol barrier” pipette tips to prepare your master mix and to aliquot the DNA. Setting up the PCR reaction with primers 8F and 1492R: PCR reagents for each group have been aliquoted and stored in the freezer. Remove the reagents and allow them to thaw on ice. See the table below for the required PCR reagents and the amounts required for a 25 µl reaction. Use the far right column to calculate the amount of reagent that you will need to put in the Master Mix. The Master Mix contains all of the reagents needed for the number of PCRs you will carry out, 24 µl of which will then be aliquoted into PCR microtubes and finally one microliter of your extracted DNA will be added for a 25 µl total reaction. Note that extra magnesium is included in the recipe, considering that it appeared to enhance the PCR run in Lab 3.

16S rRNA gene ~ 1500 bp

341F 533R

5’

5’

3’

3’

8F 1492R

Variable V3 region ~ 200 bp

Reagent Amount needed per

25 µµµµl reaction “Master Mix”: Amount

needed for N+1 Reactions (calculate)

Purified Water 11.9 µl 10X buffer 2.5 µl 5X buffer 5 µl Mg+2 1.5 µl dNTP (10 mM) 0.5 µl 5 µM Primer 8F 1 µl 5 µM Primer 1492R 1 µl Formamide 0.25 µl Taq Polymerase 0.35 µl Totals 24 µµµµl

You will be carrying out four PCR reactions: two dilutions for one of the DNA extracts, one dilution for the second DNA extract, and one negative control. You need to prepare extra Master Mix in order to account for losses and pipetting error. Therefore, prepare enough Master Mix for N+1 (5) reactions. After you have finished the calculations, and the PCR reagents have thawed, you may begin preparing the Master Mix. First, vortex all of the reagents in order to eliminate any concentration gradients which may have formed during freezing/thawing of the reagents. Then, add the reagents to a microcentrifuge tube in the order they appear on the table. Label the microtubes that you will use for the PCR reactions. Aliquot 24 µl of Master Mix into each of the PCR tubes. Finally, add 1 µl of the diluted DNA extract to each of the three sample tubes. To the negative control, add 1 µl of purified water. When all of the groups are ready- place the tubes in the thermal cycler. Until then, maintain the microtubes on ice. All of the samples will be run on the thermal cycler together- the program takes about 1.5 hours. Thermocycler program for Primers 8F 1492R:

94 ºC 2 minutes Initial denaturing step 94 ºC 30 seconds denaturing 50 ºC 30 seconds Primer annealing 72 ºC 30 seconds extension Repeat step 2-4 for 35 cycles 68 ºC 10 minute Final extension step 4 ºC � hold

Making the agarose gel:

While you are waiting for the PCR reaction to finish prepare an agarose gel. The TA will have the 1.2% agarose already melted. You will have to set up the gel casting tray and pour the mixture. Once the PCR reaction is finished you will check to see if you have products of the expected size (~1500 bp). PCR on DGGE bands: While your first PCR reaction is running, you will set up a second PCR reaction to amplify the DNA present in the DGGE bands that you cut from the gel last week. Follow the following table to prepare the master mix:

Reagent Amount needed per 25 µµµµl reaction

“Master Mix”: Amount needed for N+1 Reactions

(calculate) Purified Water 11.9 µl 10X buffer 2.5 µl 5X buffer 5 µl Mg+2 1 µl dNTP (10 mM) 0.5 µl 5 µM Primer I341f 1.25 µl 5 µM Primer I533r 1.25 µl Formamide 0.25 µl Taq Polymerase 0.35 µl Totals 24 µµµµl

This second PCR should be ready to put on the thermal cycler about the same time that the first PCR is finishing. These will be run overnight by the TA. You will check for the presence of PCR products in next week’s lab.

Lab 6: “Shot-Gun” Cloning Overview: Shot-gun Cloning is one method of obtaining 16S rRNA gene sequence information from the microbes present in your sample. You already did a PCR of the near full-length 16S rRNA genes present in your DNA extract using primers 8F and 1492R. Now you will ligate (or attach) these PCR products into a plasmid vector. A plasmid is a small circular piece of double stranded DNA external to the bacterial chromosome. Their small size makes them ideal as vectors (i.e. a carrier) for transporting DNA with genes of interest inside a cell. We will use the 4-TOPO plasmid (see Fig. 1) provided by Invitrogen for delivering your PCR product into the E. coli cells. These cells have been treated chemically so that they take up DNA readily from their environment (i.e. to make them “competent”). The process by which a competent cell takes up DNA from the environment is called transformation. Once the cells are transformed with a plasmid containing a PCR product insert, they are then spread out on Petri dishes. When spread properly, one colony originates from one cell which was transformed with a single plasmid with a single insert. Next week- you will learn how to do PCR on these individual colonies in order to retrieve the insert for sequencing.

Fig. 1: Map of 4-TOPO Cloning Vector Cloning procedure: Ligation: You will now insert your PCR products into plasmid vectors, which will then be inserted into competent E. coli cells for cloning. The “T” overhangs on the vector in Fig. 1 facilitate the ligation (or attachment) of your PCR products, which have “A” overhangs (an artifact introduced by Taq polymerase during PCR).

1.) Take 4 microliters of your PCR product into a 200 µl microtube. 2.) Add 1 µl of salt solution and 1 µl of 4-TOPO vector. 3.) Incubate the ligation reaction at room temperature for 20 minutes.

Transformation of Clones: Careful! The competent cells have been chemically treated in order to weaken their cell walls so that they can better take up DNA from the environment. Thus, they are very fragile and must be handled with care. The competent cells have been stored at -80 oC and must thaw on ice to avoid shock. 1.) Thaw the cells on ice just prior to use. 2.) Take 4 µl of your ligation reaction and add it very gently to the competent cells and stir gently with the pipet tip to mix (do NOT pipet up and down). 3.) Incubate the competent cells on ice for 20 minutes. 4.) Heat shock the cells for 30 seconds at 42 oC and place back on ice for 5 minutes. During this incubation, warm the SOC medium to room temperature. 5.) Gently add 50 µl of the SOC medium to the competent cells. Incubate at 37 °C for 1 hour in the hybridization incubator, which will turn the samples gently during incubation (Go to Purification of PCR product from DGGE bands) 6.) Plate 100 µl of the competent cells onto a prewarmed (at 37 oC) Petri dish containing LB agar and kanamycin. Use a pipeter to transfer the transformed cells onto the medium and use a sterile hockey stick to spread the cells until the liquid is absorbed into the plate. The kanamycin antibiotic will ensure that only cells that have taken up the 4-TOPO vector, which contains a kanamycin resistance gene, will grow. In order to ensure that only cells which contain a vector WITH a PCR product insert, the 4-TOPO vector contains a “gene-killer”. This means that the insert takes place within the gene-killer gene, thus inhibiting its lethal function. Therefore, if the insert is present, then this gene does not function, and the cell survives and can later be selected. The clones will be allowed to grow overnight and then the TA will store them in the refrigerator until next week when we will screen clones and prepare them for DNA sequencing. Purification of PCR product from DGGE bands. During the incubation of the competent cells you will purify the PCR products from the DGGE bands using the QIAquick PCR Purification Kit (QIAGEN). The TA already checked the presence of product in an agarose gel.

DNA Purification Procedure

1. In a 2 ml tube with a QIAquick spin column: add 100µl of Buffer PB and 20 µl of the PCR sample and mix.

2. Centrifuge at 13,000 rpm (~17,900 x g) for 60 s in a conventional tabletop microcentrifuge.

3. Discard flow-through. Place the QIAquick column back into the same tube.

4. To wash, add 0.75 ml Buffer PE to the QIAquick column and centrifuge at 13,000 rpm for 60 s.

5. Discard flow-through and place the QIAquick column back in the same tube. Centrifuge the column at 13,000 rpm for an additional 1 min.

IMPORTANT: Residual ethanol from Buffer PE will not be completely removed unless the flow-through is discarded before this additional centrifugation.

6. Place QIAquick column in a clean 1.5 ml microcentrifuge tube.

7. DNA elution: add 30 µl elution buffer to the center of the QIAquick membrane, let the column stand for 1 min, and then centrifuge at 13,000 rpm for 1 min..

IMPORTANT: Ensure that the elution buffer is dispensed directly onto the QIAquick

membrane for complete elution of bound DNA. The average eluate volume is 28 µl from 30

µl elution buffer.

You will need to run the purified product on a gel in order to estimate the concentration. Prepare three different dilutions of your purified DNA and load them in the agarose gel. Load 2 µl of each dilution and 2 µl of low mass ladder. We will document the gel on the imager and compare the intensity of the bands with the corresponding band from the ladder to determine the DNA concentration. This process will be demonstrated to you. Fill out the attached form which is used for submitting samples for sequencing to the C.S.U. macromolecular resource facility. You need to fill out the sample name and the sample concentration. The form also asks for details on the primer to be used for sequencing. To sequence the DGGE bands we need to provide a primer. We will use the primer I533r, which has a melting temperature of 47 ºC and which we will provide at 3.2 pmol/microliter.

Lab 7: PCR of Clones for sequencing and CE-SSCP Overview: You will now retrieve the inserts of your clones for DNA sequencing and identification using primers specific to the plasmid vector. You will use primers M13F and M13R (see map of vector from previous lab for location of priming sites). Because these primers are specific to the cloned vector and are not “universal” primers, then contamination is not as much of a concern as with previous PCRs using universal primers. All the same guidelines apply for PCR- vortex all of the reagents before using them, and maintain everything on ice as much as possible. PCR of clones You will prepare a PCR reaction for 10 clones plus a blank. Thus you will prepare enough Master Mix for N+1=12.

Reagent Amount needed per 25 µµµµl reaction.

“Master Mix”: Amount needed for N+1 Reactions (calculate)

Purified Water 11.4 µl 10X buffer 2.5 µl 5X buffer 5 µl Mg+2 4.5 µl dNTP (10 mM) 0.5 µl Primer M13F (20 µM) 0.25 µl Primer M13R (20 µM) 0.25 µl Formamide 0.25 µl Taq Polymerase 0.35 µl Totals 25 µµµµl

Mix all of the above in a microcentrifuge tube- vortex, and place on ice. Choose 10 colonies and circle them on the underside of the petri dish and label them 1-10. Choose colonies that are medium to large in size, and that are well-separated. Do not choose any tiny “pinpoint” colonies that may be present present, these usually grow in regions where the antibiotic has degraded and do not contain the vector with the resistance gene. (See Figure 1) You will then aliquot out 25 µl of the master mix into 11 PCR tubes using pipette tips with a white barrier. Then, lightly dip a pipette tip into one of the colonies and dip and swirl it into a PCR tube containing the master mix. Do this for all ten colonies you have chosen, using a fresh tip each time. Do not do this with the 11th tube- this is your negative control. For the negative control, just dip and swirl a clean pipette tip. After completing this, close the tubes and label them accordingly.

You will place the tubes on the thermal cycler, which will run with the following program:

94 oC 3 min 94 oC 30 sec 55 oC 30 sec 72 oC 45 sec

35 cycles

72 oC 7 min 4 oC �

This program takes about 2.5 hours.

PCR on M13 PCR products for CE-SSCP While the first PCR is running you will prepare a master mix for a second PCR. We will use the products of this PCR to do capillary electrophoresis single stranded conformation polymorphism (CE-SSCP). You will use the products of the first PCR you set up in lab today and the DNA that you extracted the first lab as templates. In this master mix the reverse primer (w104R) is labeled with a fluorescent dye (6-FAM) so that the genetic analyzer can detect it. The primers amplify the highly variable V3 region of the 16S gene and produce a product about 200 bp long (same region that was analyzed in DGGE). You will prepare a PCR for the ten M13 PCR products, your three DNA dilutions and a blank. Thus you will prepare enough Master Mix for N+1=15. .

Pick these Avoid these

Fig. 1: Tips for choosing clones.

Constituent

Amount needed per 19 µµµµl reaction

“Master Mix”: Amount needed for N+1 Reactions (calculate)

Sterile Water 11.4 µl 10x Buffer 2.0 µl Mg+2 1.0 µl dNTP [10 mM] 0.4 µl w49F [5 uM] 2.0 µl w104R [5 µM] 2.0 µl Taq 0.2 µl Total Volume 19.0 µµµµl

1. Aliquot 19 µl of the master mix into 0.2 mL tubes in the PCR. Place the tubes on ice. 2. Add 1µl of PCR product or extracted DNA to each tube. 3. Place the PCR tubes on the cycler pre-heated to 94ºC 4. Run the following program:

94 oC 2 min 94 oC 30 sec 61 oC 30 sec 72 oC 30 sec

25 cycles

72 oC 10 min 4 oC �

Running an agarose gel While the second PCR is running you will run an agarose gel with the products of the first PCR. To do this follow the steps in the protocol for lab #2. Though you will have already used these products as template for your second PCR, it will help troubleshoot if there are any problems with the second PCR.

Lab 8: Preparetion for sequencing and CE-SSCP Overview: You will purify and quantify the PCR products from your clones and DGGE bands to send them for sequencing. The purification consists on removing PCR reagents and primer dimer that otherwise would interfere with the sequencing procedure. You will also quantify the PCR product obtained with primers W49f and w104r and calculate how much you need to dilute them to use them for CE-SSCP. Purification of PCR product from DGGE bands and clones. The TA already checked the presence of product in an agarose gel.

DNA Purification Procedure

8. In a 2 ml tube with a QIAquick spin column: add 100µl of Buffer PB and 20 µl of the PCR sample and mix.

9. Centrifuge at 13,000 rpm (~17,900 x g) for 60 s in a conventional tabletop microcentrifuge.

10. Discard flow-through. Place the QIAquick column back into the same tube.

11. To wash, add 0.75 ml Buffer PE to the QIAquick column and centrifuge at 13,000 rpm for 60 s.

12. Discard flow-through and place the QIAquick column back in the same tube. Centrifuge the column at 13,000 rpm for an additional 1 min.

IMPORTANT: Residual ethanol from Buffer PE will not be completely removed unless the flow-through is discarded before this additional centrifugation.

13. Place QIAquick column in a clean 1.5 ml microcentrifuge tube.

14. DNA elution: add 30 µl elution buffer to the center of the QIAquick membrane, let the column stand for 1 min, and then centrifuge at 13,000 rpm for 1 min..

IMPORTANT: Ensure that the elution buffer is dispensed directly onto the QIAquick

membrane for complete elution of bound DNA. The average eluate volume is 28 µl from 30

µl elution buffer.

Checking purified products and CE-SSCP PCR products in agarose gel.

You will run the purified products and CE-SSCP PCR products (the TA already checked the presence of product in an agarose gel) on an agarose gel in order to estimate the concentration.

Load 2 µl of each product and 2 µl of low mass ladder with 1 µl of blue dye and 0.3 µl of the SybrGreen-DSMO solution. It is better if you mix in a 0.5 ml tube the blue dye and Sybrgreen that you need for all you samples and ladder plus 3 extra and then aliquot 1.3 µl dots in parafilm. We will document the gel on the imager and compare the intensity of the bands with the corresponding band from the ladder to determine the DNA concentration. This process will be demonstrated to you. For the purified PCR products: fill out the attached form which is used for submitting samples

for sequencing to the C.S.U. macromolecular resource facility. You need to fill out the sample

name and the sample concentration. The form also asks for details on the primer to be used for

sequencing. For the clones the primer is “T7”. You do not need to fill out any more information

than this because T7 is a standard sequencing primer and the facility will provide it free of

charge. For the DGGE bands, however, we need to provide a primer (since it was not inserted

into a cloning vector). We will use the primer I533r, which has a melting temperature of 47 ºC

and which we will provide at 3.2 pmol/microliter.

For the CE-SSCP products: calculate how much you need to dilute the products so the final concentration is about 0.6 ng/mL (prepare ~3 µl). The dilutions will be prepared in sterile water. Dilutions in water are stable only for about a week we won’t prepare them until next week.

Lab 9: Capillary electrophoresis-single stranded conformation polymorphism (CE-SSCP)

Overview: We will use CE-SSCP to analyze the PCR products of your SSCP PCR. These PCR products are fluorescently labeled (remember that the reverse primer that you used had a fluorescent dye). The principle behind SSCP is that under non-denaturing conditions, single strand DNA fragments have a folded 3D structure that is specific to the nucleotide sequence of the DNA. You will run the products through an ABI Prism 310 Genetic Analyzer (Figure 1). Fluorescently labeled samples are placed in the autosampler and injected in a capillary which is filled with a polymer. The samples are electrophoretically separated as they travel through the polymer in the capillary. As the DNA fragments pass through the window of the capillary, an argon-ion laser excites the attached dye label and they fluoresce. The analyzer detects the fluorescent signal and the data collection software converts it into an eletropherogram (it looks like a chromatogram). The mobility of the fragments is measured relative to a size standard. Ideally, each peak in the electropherogram corresponds to a single microorganism and the height of the peaks is an indication of abundance (but keep in mind that CE-SSCP is a qualitative technique or semi-quantitative in the best case). Today, you will prepare your samples for CE-SSCP. The TA and her assistant will demonstrate how to prepare the instrument for the run and start the run.

Figure 1: ABI Prism Genetic Analyzer (infohost.nmt.edu/~biology/sequencer.htm)

Autosampler region

Detection region (laser)

Capillary Gel block

region

ABI Prism® 310 Data

Collection Software

Sample pre-treatment We need to pre-treat the samples in order to obtain single-stranded DNA

1. You already calculated the necessary dilutions of the PCR product from your clones and genomic DNA. Now, you are going to prepare these dilutions in freezer tubes. Remember that you need ~0.5 ng/ul of PCR product from clones and ~ 0.7 ng/ul of PCR product from your community DNA.

2. Prepare a master mix using Hi-Di Formamide and the ROX size standard for the number

of samples plus one blank. The Hi-Di Formamide cannot be defrosted more than once, so any left over formamide must be thrown away. Combine the needed amount of each reagent in a 1.5mL tube and vortex to ensure homogeneity.

Master Mix for

1 Sample Master mix for all samples

Component Volume [�L] HiDi Formamide 18.8 ROX size standard 0.2

3. Assemble the sample tray (demonstrated in class). 4. Aliquot out 19uL of your master mix into each tube and add 1uL of your sample into the

appropriate tube.

5. Push the septa strips into the tubes so that each tube is covered.

6. Leaving the septa strips on, denature your samples for 5 minutes at 95 °C in a thermocycler. Immediately place you samples into ice water for 15 minutes and keep refrigerated until ready to use. This causes the single DNA strands to fold up onto themselves (rather than form double strands), forming the 3-D shapes by which they will be separated.

7. Remove any air bubbles from the sample tubes by gently tapping the outside of the tube

or by spinning the samples in the salad spinner. The entire sample tray can be placed in the salad spinner.

8. When you are ready to begin a run, place a retainer clip directly over each septa strip.

The retainer clip will snap onto the sides of the sample tray and will fit into the holes of the septa strips. Make sure the retainer clips are pushed down as far as they can go and that they are all level with each other to keep the electrode from bending.

9. Move the autosampler tray forward by pushing the autosampler button located on the left

side of the instrument. Place the sample tray on the autosampler. The PCR tube rack is no longer needed.

10. Return the autosampler to its original position, close the instrument doors, and begin your run (The TA will demonstrate how to do this).

Lab 10: Nuts and Bolts of DNA Sequence Analysis

Overview: You will now analyze your sequences from both the DGGE bands and the clones to determine the identity of the organisms represented by these sequences. Sequence analysis for the purpose of classifying organisms, or phylogenetics, is a field in and of itself, and we will only be able to scratch the surface with this exercise. The purpose of this lab, therefore, is to give a basic idea of the techniques involved and allow you to identify your microorganisms. For further interest, see Phylogenetic Trees Made Easy: A How To Manual for Molecular Biologists by Barry Hall. Sequence File format: Each group will be given a group of sequence files, including chromatogram files with “.ab1” file extension. This chromatogram file can be opened by a program called Chromas, which is a free software available on the internet at: http://www.technelysium.com.au/ . This program will already be downloaded on the computer. Open each file and look at the overall quality of the chromatograms. Note that each base is color coded in the chromatogram (A is green, C is blue, G is black, and T is red). The sequence reader judges the identity of the nucleotide at each position by comparing the relative heights of the peaks. If two peaks are overlapped, then the program is not able to judge what the nucleotide is, and you will se a pink “N” (unknown) in that position. The following figures show examples of high quality sequence data, and poor quality sequence data, as viewed in Chromas.

Fig. 1: Example of a chromatogram with good quality sequence data.

Fig. 2: Example of a chromatogram with poor quality sequence data.

Sequence Analysis: If you have high quality sequence data- then the analysis will be simple- if not…….it will be challenging. The most common reason that a chromatogram has a noisy signal is that there were multiple templates present during the sequencing reaction. Considering that our DGGE bands did not resolve very well, it is likely that more than one sequence was present in some of the bands which you cut. If you sequence multiple sequences at the same time- then the signals will overlap and you will get a noisy signal. The clones are more likely to give a cleaner signal because they were well resolved as individual colonies. After you open each file and take note of the sequence quality you will begin to analyze the sequences. Go back to the beginning of your sequence list and open the file in Chromas. Go to “File\export” and export the file in the “FASTA” format [alternatively, the exported text file may already be available]. FASTA is a very commonly used format for sequence files. Once you have exported this file- you can open it in MS Word. Word allows you to edit the file using the “Search\Replace” functions, etc., but save the file as “TEXT only” whenever you modify it. After you open the file in Word, copy the sequence portion of the file (everything after the first line- the first line contains the sequence name and the formatting commands for FASTA format) onto the clipboard. Now open your web browser and go to the National Institute of Health BLAST website: http://www.ncbi.nlm.nih.gov/BLAST/. This website links to the most comprehensive and up-to-date sequence information available. The advantage of Blast is that it will allow you to check your sequence against this vast database, and will give you a visual alignment of the closest matches, which can help you correct any potential errors in your sequence. Click on “nucleotide-nucleotide” Blast and paste your sequence in the search window. Then click “Blast”. After this, a new screen will come up- when this happens, click “Format”. After this Blast will begin to query the database. This could take a few seconds, or several minutes, depending on the traffic to the website at the time of your query. After some time, your alignment will come up. SAVE the alignment, you will have to open it again when you create a phylogenetic tree. When this happens, you will see something like Fig. 4. The top portion is a summary of your sequence matches. Red color indicates a good match along the length of your sequence, while pink indicates that there is a lower match along the length of your sequences. If you see green, blue, or black, then that indicates that your sequence is most likely too poor to analyze any further. Scroll down and look at the names of the organisms which gave the closest matches. Note that it is not uncommon that several of these will be “unknown” or “uncultured” bacteria. Remember that most bacterial have not yet been cultured, and the only thing we know about them is their sequence information, so this should not be surprising. Also note that this format does not tell you much about how to classify your sequence- we will use a different website- the Ribosomal Database Project- to do this. In some cases, clicking on the “taxonomy report” link in Blast can help provide a preliminary classification. Now scroll down and look at the alignments. This gives you a chance to edit your sequence and look for and repair errors. This is a tricky business, and you must do this very CONSERVATIVELY. The following are some general guidelines for sequence editing:

Distribution of 100 Blast Hits on the Query Sequence

Mouse-over to show defline and scores. Click to show alignments

Sequences producing significant alignments: (bits) Value gi|30103112|gb|AY177357.2| Phenanthrene-degrading bacterium... 337 7e-90 gi|23345134|gb|AY136080.1| Sphingomonas sp. KIN84 16S ribos... 337 7e-90 gi|4868348|gb|AF131297.1|AF131297 Sphingomonas sp. JSS-54 1... 337 7e-90 gi|12247763|gb|AF327069.1|AF327069 Sphingomonas sp. SA-3 16... 337 7e-90 gi|456233|dbj|D13727.1|SPP16SRR6 Sphingomonas terrae gene f... 337 7e-90 gi|30103121|gb|AY177366.2| Sphingomonas sp. 86 16S ribosoma... 329 2e-87 gi|30060219|gb|AY254693.1| Uncultured alpha proteobacterium... 329 2e-87 gi|40240919|emb|AJ619081.1| uncultured alpha proteobacteriu... 329 2e-87 gi|19699044|gb|AY081981.1| Uncultured bacterium clone KRA30... 329 2e-87 >gi|30103112|gb|AY177357.2| Phenanthrene-degrading bacterium M20 16S ribosomal RNA gene, partial sequence Length = 1357 Score = 337 bits (170), Expect = 4e-90 Identities = 170/170 (100%) Strand = Plus / Plus Query: 1 cctacgggaggcagcagtggggaatattggacaatgggcgaaagcctgatccagcaatgc 60 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 264 cctacgggaggcagcagtggggaatattggacaatgggcgaaagcctgatccagcaatgc 323 Query: 61 cgcgtgagtgatgaaggccctagggttgtaaagctcttttacccgggatgataatgacag 120 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 324 cgcgtgagtgatgaaggccctagggttgtaaagctcttttacccgggatgataatgacag 383 Query: 121 taccgggagaataagctccggctaacttcgtgccagcagccgcggtaata 170 |||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 384 taccgggagaataagctccggctaacttcgtgccagcagccgcggtaata 433

Fig. 3: Example of Blast Alignment.

Guidelines for Sequence Editing: Scroll down the alignments and look for errors. The most problematic yet easy to fix error is a “Gap”. A gap occurs when the DNA sequencer either inserts an extra base- or removes a base- this causes a shift in the alignment. Figure 3 presents a “perfect” or 100% match- and will not need any further editing. Figure 4 presents an example of an alignment with gaps which need to be edited. If gaps are found in Blast- then go back to the Chromas file and find these positions on the chromatogram (the numbers in both Blast and Chromas can help guide you). You will most likely notice that in the chromatogram at this position that it is either missing a base pair or one has been added. Insert/Delete this base pair as appropriate. This is considered to be a conservative repair because gaps between closely related species are evolutionarily unlikely. Figure 5 shows the corresponding chromatogram for Figure 4. Can you find the gaps? gi|22002633|gb|AY122605.1| Uncultured bacterium clone OSS-41 16S ribosomal RNA gene, partial sequence Length = 583 Score = 333 bits (168), Expect = 1e-88 Identities = 191/196 (97%), Gaps = 2/196 (1%) Strand = Plus / Minus Query: 30 tattaccgcggnctgctggncacgtagttagccggtgcttattcttacggtaccgtcatg 89 ||||||||||| ||||||| |||||||||||||||||||||||||||||||||||||||| Sbjct: 195 tattaccgcgg-ctgctgg-cacgtagttagccggtgcttattcttacggtaccgtcatg 138 Query: 90 tgccccaggtattaaccagagccttttcgttccgtacaaaagcagtttacaacccgaagg 149 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 137 tgccccaggtattaaccagagccttttcgttccgtacaaaagcagtttacaacccgaagg 78 Query: 150 ccttcttcctgcacgcggcattgctggatcagggttgcccccactgtccaaaattcctca 209 |||||||||||||||||||||||| |||||||||||||||||| ||||||||||||| || Sbjct: 77 ccttcttcctgcacgcggcattgcaggatcagggttgcccccattgtccaaaattcccca 18 Query: 210 ctgctgcctcccgtag 225 |||||||||||||||| Sbjct: 17 ctgctgcctcccgtag 2

Fig. 4: Example of an aligned sequence with gaps.

Fig. 5: Chromatogram corresponding to Figure 4 with gaps, in this case, two “N”s have been inserted where they should not have been. Editing “N”s Other sequence editing, such as editing Ns is less conservative than editing gaps, and should be done with caution. One rule of thumb: Check at least 10 alignments for the corresponding nucleotide present at the position of an N for the most closely related species. If for example, 10 out of 10 of these show a “G”, go back to your chromatogram and look at this position. If G is indeed the highest peak at this position- then you may change it- however you should maintain any changes in lower case. What if not all of the alignments show the same base pair at this position, or if the peak corresponding to that base pair is not the highest? It is best to maintain that position as an N. Another option that Fasta format options is for you to narrow this down using alternative symbols which represent more than one base pair. See Figure 6: Export your corrected chromatogram, open the file in Word. Once you have your sequence edited- save the file. You should also save the html file with the Blast matches- this can be helpful later on if you need to go back to it, and you do not have to wait to query the database again. Go to File, Save as, and save as html (default). You may get an error message (ignore). A --> adenosine M --> A C (amino) C --> cytidine S --> G C (strong) G --> guanine W --> A T (weak) T --> thymidine B --> G T C U --> uridine D --> G A T R --> G A (purine) H --> A C T Y --> T C (pyrimidine) V --> G C A K --> G T (keto) N --> A G C T (any) - gap of indeterminate length Figure 6: Summary of FASTA format symbols. Using the Ribosomal Database Project to Classify Your Sequence You may have noticed that Blast was not very helpful for classifying the organism represented by your sequence. For this we will use the Ribosomal Database Project (RDP). This website is maintained by the Center for Microbial Ecology at Michigan State University, and is an effort to phylogenetically classify all known microbial sequences.

Open your corrected chromatogram file in Word. You want to use this corrected sequence for RDP because it does not allow you to view and correct the actual sequence. Go to http://rdp.cme.msu.edu/ and click on “Sequence match”. After a few seconds, a tree should come up which shows the current classification of the sequences most closely related to yours. This database is not as updated as Blast- so your matches may not be as high, but you should have a better idea of how to classify your sequence. You may also save the RDP html file for future reference. Figure 7 shows an example RDP output. You can click on each level of the taxonomy to find more information on related microorganisms. Lineage (click node to return it to hierarchy view): Hierarchy View: [ options ]

domain Bacteria (1) (query sequences) phylum Proteobacteria (1) class Gammaproteobacteria (1) order Xanthomonadales (1) family Xanthomonadaceae (1) genus Stenotrophomonas (1) unknown [view selectable matches]

Data Set Options:

Strain: Type Non Type Both Source: Uncultured Isolates Both

Size: >1200 <1200 Both KNN matches:

Refresh

Strain: View only sequences from species type strains, non-type strain sequences or both. Type strain information is provided by Bergey's Trust. Hint: Type strains link taxonomy with phylogeny. Include type strain sequences in your analysis to provide documented landmarks.

Source: View only environmental (uncultured) sequences, only sequences from individual isolates, or both. Source classification is based on sequence annotation and the NCBI taxonomy.

Size: View only near-full-length sequences (>1200 bases), short partials, or both.

Fig. 7: Example of a Ribosomal Database Project Sequence Match output.

Congratulations! You should now be able to identify your sequence at least to the Phylum or Class level, and with some luck, possibly to genus.

Finally, fill out a Table like the following in your lab book so that you can keep record of your sequences (The first one is done for you as an example). The last column is where you can make comments about the quality of the sequence data, any gaps removed, or Ns repaired, etc. The number of bp match, and percent match can be determined from the Blast Alignment.

Seq Name Blast ID

% match

# bp match

Ribosomal Database ID Comments

DGGE band 2ea1-5

Alpha Proteobacteria, Sphingomonas (phenanthrene degrading)

99% 170 bp

Alpha Proteobacteria, Sphingopyxis/Sphingomonas genus

Excellent Sequence Data

Clone #1

Constructing a Phylogenetic Tree Using DNA Sequences Overview: A phylogenetic tree is used to represent the relationship between groups of organisms. Historically, these relationships were based on morphology. However, molecular biology has allowed phylogenetic trees to be constructed showing the similarity of molecular sequences such as DNA or proteins. These trees are seen as a tool for understanding biological processes and their relationships to other sequences of interest. Creating a Phylogenetic Tree Choosing reference sequences.

1. Open the BLAST alignments of all your sequences. Look for microorganisms that appear frequently in the alignment or for microorganisms that you would expect to find in your samples. You are going to chose five of these sequences and use them as reference to compare with your sequences.

2. To chose a sequence: a. In the alignment list check the box that is next to the “g-number” of the sequence

you want to download. For example:

> gi|7595962|gb|AF251436.1| Ferrimicrobium acidiphilum 16S ribosomal RNA gene, partial sequence Length=1449

b. After you selected the sequence/s scroll down to the bottom of the list. Click on

“Get selected Sequences”. The next screen will look like this:

c. In the Display option choose FASTA and then go to Send To and choose file. A message box will open. Save the files.

Creating an input file for ClustalX.

1. We will do this by creating a text file with all your sequences and the reference sequences that you selected.

2. To create a text file: a. Open your edited sequences (they were saved in FASTA format) in MSWord. b. Open a new file in MSWord or Notepad and copy your sequences to this new file.

NOTE: ClustalX interprets > as the start of a line. All characters after > up to the first space are part of the file name. (File names should be less than 10 characters, and include no spaces or special characters other than the underscore.) All characters after the first space are considered the sequence. An input file should look like the box below. Use copy and paste commands to insert your sequences into one file.

c. File � Save As Filename.txt (Save as text file type)

Creating an alignment in ClustalX 1. File � Load Sequences and load the input file you just created

>sequencename1 ctgagctaccggttaacca… >sequencename2 accggttaaccctgagcta… >sequencename3 gttaaccctagtctgagcta…

2. Alignment � Output format options and select Clustal format, Nexus format, and ClustalW Sequence Numbers on. Click Close.

3. Alignment � Do complete alignment � Align 4. Below the alignment is a histogram that indicates the degree of similarity. (Peaks

indicate high similarity; valleys indicate low similarity.) The * above the sequences indicates positions that have been fully conserved.

Trimming the sequences 1. All the sequences in your alignment should be the same length for proper comparison.

Any longer sequences should be truncated so you are comparing sequences of the same length.

2. Open Proseq2 3. File � Open � Files of type: Fasta 4. This allows you to see all the files in the folder 5. Select filename.nxs and change Files of type: Nexus. Open the nexus file created from

the alignment. 6. Trim the sequences to the same size by selecting the part you want to cut and go to Edit � Cut

7. File � Save As: filename.nbr to save the trimmed sequences.

Creating a phylogenetic tree in Clustal. 1. Open filename.nrb in Clustal 2. Trees � Bootstrap N-J Tree. It will open a message box indicating where the tree will

be saved. Write this down so you know where to look for your file.

Viewing the tree in TreeViewX 1. Start TreeVew 2. File � Open �open the *.phb file. 3. Tree formats

a. Cladogram: shows only the branching order of nodes. Can be presented in a rectangular or slanted format.

b. Phylogram: shows branching order and distance information c. Format text by selecting Trees � Leaf font and choosing your font

settings 4. Saving trees

a. Edit � Copy allows you to copy the entire tree. If the tree is pasted in a program such as power point that allows you to “draw,” you can convert the tree to a Microsoft picture, ungroup the object, and edit the tree.

b. File � Save As Picture allows you to save the tree as a picture that can be imported into documents.

Lab 11: Real-Time PCR-Part I

Overview: Real-time PCR (also quantitative-PCR or QPCR) is a quantitative version of PCR that uses a fluorescent signal to monitor product formation in “real time”. The Ct value, or threshold cycle, correlates with the amount of template originally present. A low Ct value indicates a high initial concentration of target DNA and vice versa. We will use QPCR to quantify total bacteria (part I) and specific groups of microorganisms that you might be interested in studying in your sample such as sulfate-reducers, ammonia-oxidizing bacteria, or antibiotic-resistant microorganisms (part II). We will meet at ERC for this lab. Real-time TaqMan PCR to Quantify Total Bacteria:

We will use a TaqMan PCR assay to quantify the total bacterial populations present in your samples. We will use universal PCR primers 1369F and 1492R to amplify a 123 bp region of the 16S gene. We will also use probe TM1389F, which targets a region between the two primers and carries the fluorescent dye Cy-6 on the 5’ end and the black hole quencher (BHQ) on the 3’ end (Fig. 1). As the Taq enzyme amplifies the region between the primers, it encounters the TaqMan probe and the 5’ -3’ exonuclease activity of the enzyme releases the dye end of the primer from the quencher, and thus releases the fluorescent signal. The fluorescent signal is detected by the real time cycler and the data is collected by the Cephid SmartCycler software. The tubes used for the SmartCycler are specialized to maximize the signal intensity (Fig. 2). Setting up the Reactions: Each group will set up duplicate reactions for two DNA extractions. The Smart Cycler can process 16 tubes at the same time. Each group is going to have a total of 5 tubes (two samples in duplicate + one blank). Therefore, we are going to run the samples in three runs. Each group will make a Master Mix for their samples according to Table 1. Prepare enough for N+1 reactions (4 samples, 1 blank +1 = 6 reactions per group).

After you have prepared the Master Mix, aliquot 48 microliters into two microtubes. Add 2 microliters of your first DNA extraction to the first tube, and 2 microliters of the second DNA extraction to the second tube. Mix each tube thoroughly by vortexing, then transfer 25 microliters of each mix into two Smart Cycler tubes for each sample. Preparing the tubes in this

1369F TM1389F

5’

5’

3’

3’

1492R

Fig. 1: Schematic of TaqMan PCR of Total Bacterial 16S rRNA gene

BHQ Cy-6

way helps to get better duplication of the results. Transfer the remaining master Mix to the 5th Smart Cycler tube for the blank.

Fig. 2: Schematic of Cepheid SmartCycler System

Table 1: Setting up the Real Time PCR Reaction.

Running the Samples: After you have prepared the SmartCycler tubes, maintain them on ice until all groups are ready to run the samples. When all groups are ready, transfer them to the cycler- they will be run with the following program: 95 ºC 2 minutes Initial denaturing step 95 ºC 15 seconds denaturing 53 ºC 60 seconds Primer annealing 72 ºC 20 seconds extension Repeat step 2-4 for 50 cycles Smart Cycler cannot be set a temperature lower than 45 ºC

The run will require 1.5 hours. After the run is finished, determine the Ct value for each sample. To convert the Ct value, use the following formula based on the previous calibration: Log [DNA(ng/ul)] = -0.2591 Ct + 3.4253 Compare the value that you obtain with the original concentration of DNA that you had calculated based on absorbance at 260 nm.

Reagent Amount needed per 25 µµµµl reaction

“Master Mix”: Amount needed for N+1 Reactions

(calculate) Purified Water 13.28 µl 10X buffer 2.5 µl 5X buffer 5.0 µl dNTP (10 mM) 0.5 µl Primer 1369F 0.25 µl Primer 1492R 0.25 µl TaqMan Probe 0.37 µl Taq Polymerase 0.35 µl Mg 2+ 1.5 µl

Total 24 µµµµl

Lab 12: Real-Time PCR-Part II

Overview: In the previous lab you quantified total bacteria in your samples using real-time PCR. Now, we will look at specific microorganisms that might be of interest in your samples such as ammonia-oxidizing and sulfate-reducing bacteria (AOB and SRB respectively). To quantify AOB we will use a TaqMan PCR. For SRB we will use a SYBRGreen I PCR. SYBRGreen I is a DNA-binding dye that incorporates into double stranded DNA. It has an undetectable fluorescence when it is in its free form, but once bound to the dsDNA it fluoresces. Its greatest advantage is that it can be used with any pair of primers for any target. We will meet at ERC at 2:30 pm for this lab.

Choosing target microorganisms

AOB are strict aerobes. On the other hand most SRB are strict anaerobes (there are some SRB that are aero-tolerant). If your sample was taken from an aerobic system (eg., aerobic digester, surface water) you might want to target AOB. If your sample came from an anaerobic system (eg., groundwater, sulfate-reducing bioreactors, sediments) then you might want to target SRB. SRB are a very phylogenetically diverse group. In this lab we will target a group of SRB known as Desulfobacteria. Setting up the reactions: Each group will set up duplicate reactions for the two DNA samples used to quantify total bacteria. Each group will make a Master Mix for their samples according to Table 1(for SRB) or Table 2 (for AOB). Prepare enough for N+1 reactions (4 samples, 1 blank +1 = 6 reactions per group). After you have prepared the Master Mix, aliquot 48 microliters into two microtubes and 24 microliters into a third tube (this is for the blank). Add 2 microliters of your first DNA extraction to the first tube, and 2 microliters of the second DNA extraction to the second tube. Mix each tube thoroughly by vortexing, then transfer 25 microliters of each mix into two Smart Cycler tubes for each sample. Preparing the tubes in this way helps to get better duplication of the results. In the blank tube add 1 µl of water and then transfer 25µl to a Smart Cycler tube.

Table 1: Master Mix for SRB

Reagent Amount per 25 µµµµl reaction Amount needed for N+1 Reactions

SyBR Green master mix 12.5 µl Primer HDBM 52f 0.75 µl Primer HDBM 372r 0.75 µl H2O 10 µl Total 24 µµµµl

Running the Samples: After you have prepared the SmartCycler tubes, maintain them on ice until all groups are ready to run the samples. When all groups are ready, transfer them to the cycler- they will be run with the corresponding program:

Program for AOB 95 ºC 300 seconds Initial denaturing step 95 ºC 30 seconds denaturing 60 ºC 60 seconds Primer annealing 72 ºC 15 seconds extension Repeat step 2-4 for 45 cycles

Program for SRB 95 ºC 900 seconds Initial denaturing step 94 ºC 15 seconds denaturing 63 ºC 30 seconds Primer annealing 72 ºC 30 seconds extension Repeat step 2-4 for 45 cycles To convert the Ct value, use the following formula based on the previous calibration: Desulfobacteria: Log [DNA(ng/ul)] = -0.2966 Ct + 4.8437 AOB: Amo gene (copies/ ul) = 5 x 1011 x e^(-0.6493*Ct)

Table 2: Master Mix for AOB Reagent Amount per 25 µµµµl reaction Amount needed for N+1

Reactions Purified Water 13.28 µl 10X buffer 2.5 µl 5X buffer 5.0 µl dNTP (10 mM) 0.5 µl Primer AOB1149f (10µM) 0.25 µl Primer AOB1295R (10µM) 0.25 µl TaqMan Probe 0.37 µl Taq Polymerase 0.35 µl Mg 2+ 1.5 µl

Total 24 µµµµl

Lab 13: Fluorescent in situ Hybridization Part I. Overview: We are going to use fluorescent in situ hybridization (FISH) to look at your samples. The principle of FISH is based on the ability of single-stranded DNA to anneal to complementary RNA. Fluorescently labeled oligonucleotides (probes) specific to different types of microorganisms are used to determine the microbial composition of the sample. FISH has the advantage of not requiring nucleic acid extraction or PCR (it is not subject to PCR biases) and it also allow us to quantify the microbial groups in our sample. We will use four different probes which target total bacteria, and three classes of Proteobacteria. Since all the probes are labeled with the same fluorescent dye (fluorescein) we cannot hybridize the same sample with all the probes at the same time. Instead, we are going to prepare the samples in four replicates and each replicate is going to be hybridized with a different probe. We are going to visualize the samples next week with a Nikon E200 microscope with an epi-fluorescence attachment. Procedure: Your samples were fixed during lab 1. The cell fixation was performed using approximately 1 g of sample which was centrifuged to pellet the cells and then resuspended in 4% paraformaldehyde. After one hour this solution was centrifuged and the pellet resuspended in 1X PBS buffer. Finally, this solution was centrifuged again, the pellet resuspended in storage buffer and stored at –20oC. Hybridization and Washing

1. Label 4 microscope slides with a pen (not a Sharpie!). To label the slides remember that all the slides are going to have the same sample. The difference is going to be the probe that you are going to use in each of them. The following table shows the probes that you are going to use in this lab:

Target Probe

name Probe sequence (5’ to 3’)

All Bacteria EUB338 GCTGCCTCCCGTAGGAGT β -Proteobacteria BET42a GCCTTCCCACTTCGTTT γ-Proteobacteria GAM42a GCCTTCCCACATCGTTT δ-Proteobacteria delta402 CGGCGTCGCTGCGTCAGG

2. Pipette 10 µl of well-mixed, “fixed”environmental sample onto each of the glass

microscope slides. 3. Allow the microscope slide to dry for 5 minutes at 46oC. 4. Dehydrate the sample by submerging the microscope slides in 50%; 80%; and 96%

Ethanol for 1 min, each. 5. Allow the microscope slides to dry for 5 minutes at 46oC. 6. Pipette 9 µl of hybridization buffer onto the sample.

NOTE: The stringency required during hybridization is not the same for all the probes. The TA is going to prepare the hybridization buffers but make sure you use the right

buffer for each probe. The following table summarizes the composition of the hybridization and wash buffers for each probe:

Probe EUB 338 BET42a GAM42a delta402

Formamide [µµµµl] 400

(20%) 700

(35%) 700

(35%) 600

(30%) NaCl 5M [µµµµl] 360 360 360 360

Tris-HCl pH 7[µµµµl] 40 40 40 40 10% SDS [µµµµl] 2 2 2 2

H2O [ml] 1198 898 898 998

Hyb

ridi

zatio

n bu

ffer

Final volume[ml] 2 2 2 2 Tris-HCl pH 7[µµµµl] 1000 1000 1000 1000

NaCl 5M [µµµµl] 2150 501 501 1020 EDTA 0.5M[µµµµl] 500 500 500 500 10% SDS [µµµµl] 50 50 50 50

H2O [ml] 46.30 47.95 47.95 47.43 Was

h B

uffe

r

Final volume[ml] 50 50 50 50

7. Pipette 1 µl of fluorescently labeled oligonucleotide probe (50 ng/µl) onto the sample. 8. Place the microscope slides into a 50 ml conical tube. Place a paper towel with the

remaining hybridization solution bellow the slide. Hybridize for 3 hr at 46oC. The TA is going to wash your samples after the hybridization: 9. Prewarm 50-ml of Wash Buffer to 48oC. 10. Rinse the microscope slides with 10-ml of wash buffer to remove excess probe. 11. Wash the sample by submerging the microscope slide in 40-ml of wash buffer for 1 hr at

48oC. 12. Rinse the microscope slides with water. 13. Allow the microscope slides to air dry. 14. Store the microscope slides in the dark at –20oC for two days to one week.