CE 742 Advanced Topics in Environmental Engineering Lab Manual
Overview:
Each team has been assigned so that at least one partner has some experience with molecular
techniques. Each team should choose a sample relevant to Environmental Engineering which
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:
Ethidium Bromide (EtBr): This is used to stain gels and is a suspect carcinogen.
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 3 pipeters with 3
volumes: 0.5-10 l, 10-100 l, and 100-1000 l. 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
Luciana Pererya, Mary Beth Talty, and Chris Messersmith
Group 2
Ruoting Pei and Mustafa Yarkin
Group 3
Anurita and Matt Stephens
Group 4
Matt Hoelscher and Winnie Lin
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. Extraction will be verified by agarose gel electrophoresis and quantified by
spectrophotometry.
Procedure:
Selection of Sample:
Each team should select a sample of Environmental Engineering relevance for DNA extraction.
Examples include: bacterial cultures (relatively easy to extract), soil (relatively more difficult),
sediment, 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 per sample is required. 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. If you
are unsure, ask the instructor for advice on what sample and how much to bring.
Agarose Gel Preparation:
First you will prepare an agarose gel. You will need this after DNA extraction to verify that it
worked. You will also use this procedure throughout the semester to visualize DNA.
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.) A 2.5% solution of agarose in 1X TAE buffer (already prepared and melted at 55ºC).
*Caution! Wear gloves!:This solution contains 3 l per 100 ml of ethidium bromide, a
DNA intercolating agent, which binds to DNA and fluoresces under UV light. This will
allow you to visualize the DNA later. This agent is also a suspect carcinogen, and should
not be allowed to contact the skin.
3.) Take this solution and pour 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 pipet tip.
4.) Set aside and allow the gel to solidify.
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 pipet tip) to transfer the sample into the tubes.
2.) Add 978 microliters of phosphate buffer and 122 microliters of MT Buffer to the matrix
tube using the 100-1000 microliter pipeter.
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 l 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 pipet tip and discard, taking care not to remove any
settled binding matrix.
8.) Resuspend the sample by swirling and transfer 600 microliters 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 l 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.
11.)
Add 50 l of DES (ultrapure water, DNA and pyrogen-free) and stir the binding
matrix gently with the pipet 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! Be sure to label this final tube appropriately for storage
(Name, date, sample identirication). Check product on an agarose gel and store sample at
-20ºC.
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 (black)
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 up until the level of the buffer is flush with the top of the
gel.
B. Preparing the Samples and Loading the Gel
5.) Now cut a piece of parafilm (about 2 inch by 4 inch should be sufficient) and place on the
lab bench, parafilm side up.
6.) Using the 0.5-10 l pipeter, place fourl “dots” of blue loading dye on the parafilm
(one dot for each of the samples that were extracted, and one dot for each of the
molecular weight standards).
7.) Now take 3 l of the first molecular weight marker, and add to the first blue dot. Mix by
pipeting up and down. Once mixed, take the sample and dye mixture back up in the tip.
8.) 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.
9.) Repeat this procedure with the two DNA extracts, and finally with the second molecular
weight marker.
C. Running and Visualizing the Gel:
10.)
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 120 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 240 v.
Run until you see the blue dye move about halfway down the gel. At this time, turn off
the voltage to the gel.
11.)
In order to “see” the DNA, you need to look at it
1.
2.
under UV light. Take the gel over to the hand-held UV lamp. Make sure that you are
wearing the UV protection shield over your face, and that any bare skin is covered
completely by gloves and your sleeves. Turn on the lamp
and hold over the gel. 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 = 1500 bp, brightest band (middle) = 500 bp, and bottom
band = 100 bp. 2.) Extract of genomic DNA from E. coli.
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. 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 microliters (eg. 5
microliters of DNA extract added to 495 microliters 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 ultrapure 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 pipeter.
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 / l = x ng / l
To calculate the concentration of genomic DNA in the extract before dilution:
x ng / l * (500 l / 5 l) = y ng / l
To calculate the total mass of genomic DNA in the extract:
y ng / l * 50 l = z ng
Congratulations! You have completed your first lab! Be sure to record your
observations in detail in your lab book.
Lab 2: Polymerase Chain Reaction (PCR)
Objective:
You will use PCR to amplify the 16S rDNA genes of the bacterial community DNA which you
have extracted. We will take a “nested” PCR approach. This will require two PCR reactions: 1)
amplifying near full-length of the 16S gene (~1500 bp) using primers 8F and 1492R 2) Using the
product of the first PCR reaction as template to amplify the ~200 bp V3 region within the 16S
gene using primers 341F and 533R. The PCR product from the first PCR will be used later in
the semester for cloning, while the second PCR product will be used next week for denaturing
gradient gel electrophoresis (DGGE).
16S rRNA gene ~ 1500 bp
5’
8F
341F
F
533R
3’
3’
1492R
5’
Fig. 1: Position of PCR primers for “nested” 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
increased 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” which 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 8F and 1492R:
PCR reagents for each group have been aliquotted and stored in the freezer. Remove the
reagents and allow them to thaw on ice. For the first PCR reaction you will need:
Amount needed per 25 l
reaction
Reagent
“Master Mix”: Amount
needed for N+1 Reactions
(calculate)
13.15 l
2.5 l
5
l
1
l
1
l
1
l
0.35 l
Purified Water
10X buffer
5X buffer
dNTP (10 mM)
Primer 8F
Primer 1492R
Taq Polymerase
Totals
24 l
See the above table for the required PCR reagents and the amounts required for a 25 l 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 l of which will then be aliquotted into PCR microtubes and finally one
microliter of your extracted DNA will be added for a 25 l total reaction.
You will be carrying out 4 PCR reactions: 1 for each of the extracted DNA samples, 1 negative
control, and 1 positive 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 l tube = $300.00!). After you have added all
of the reagents- vortex and place on ice.
Based on the concentration and purity of your DNA extract, you will need to do a dilution of
your DNA before using for PCR. You should have calculated the concentration of your DNA at
the end of the last lab- also determine the ratio of absorbance at 260 nm:280 nm. If this ratio is 2
or greater, then you have relatively pure DNA and you will not have to do a high dilution (1:3 is
probably a good dilution to try in this case). If this ratio is less than 2, then try a higher dilution
(1:5, 1:10 or 1:20 for high concentration DNA). Do not dilute the whole DNA extract, insteadadd the required volume of dilution water to a microtube and add the DNA extract to this tube.
For example, for a 1:5 dilution, add 4 l of water to a microtube, and 1 l of DNA extract.
Now label the microtubes that you will use for the PCR reactions. Aliquot 24 l of Master Mix
into each of the PCR tubes. Finally, add 1 l of the diluted DNA extract to each of the two
sample tubes. To the positive control, add 1l of the positive control, and to the negative
control, add 1 l 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
94 ºC
30 seconds
50 ºC
30 seconds
72 ºC
30 seconds
Repeat step 2-4 for 24 cycles
68 ºC
10 minute
4 ºC
∞
Initial denaturing step
denaturing
Primer annealing
extension
Final extension step
hold
Making the agarose gel:
While you are waiting for the first PCR reaction to finish you will make an agarose gel. To
make a 1.2% agarose gel, weigh out 0.9 grams of agarose and transfer to a 250 ml flask. Add 1.5
ml of 50X TAE buffer and 73.5 ml (this can be approximate) of DI water. Swirl the mixture to
mix and microwave 1 minute on high. Swirl again to mix and microwave again, watching
closely. Just as the mixture begins to boil- stop the microwave. Swirl and make sure that the
mixture is clear and there is no undissolved agarose. Add 1 l of ethidium bromide and swirl
once more to mix. 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. While the gel cools,
begin setting up the second PCR reaction.
Setting up the second PCR Reaction with primers 341F and 533R :
Now make the calculations for N+1 reactions with primers 341F and 533R. You will use the
four PCR products from the first PCR as template, plus a new blank (N+1=6).
Amount needed per 25 l
reaction
Reagent
Purified Water
10X buffer
5X buffer
dNTP (10 mM)
Primer 341F
Primer 533R
Taq Polymerase
13.15l
2.5 l
5
l
1
l
1
l
1
l
0.35 l
Totals
24 l
“Master Mix”: Amount
needed for N+1 Reactions
(calculate)
Prepare the Master Mix, as you did for the first PCR. Note that the only difference will be the
primers used. Vortex the Master Mix and aliquot 24 l into the PCR microtubes. Hold on ice
until the first PCR is done
Load and Run the Agarose Gel:
Once the gel has cooled, 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.
Once the first PCR has finished, load 3 l of the PCR product onto the agarose gel, using 3 l of
blue loading dye and parafilm as you did for Lab 1. Don’t forget to load the DNA molecular
weight standard (“ladder”). Load 5 l of the standard (last week it seemed that 3 l was not
enough).
Run the gel at 150 volts 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 1500 bp.
If your reaction is successful, you should see a band at ~1500 bp (see appendix for molecular
weights of bands in size standard).
Running the Second PCR: Primers 341F and 533R:
After you have checked your PCR product from the first reaction, prepare a dilution of the PCR
product. If the product looked very strong, make a 1:5 dilution, if it looked weak, make a 1:3
dilution, if you were not able to see any product, do not make a dilution.
Add 1 l of the diluted PCR product (including the first blank) to the new PCR tubes and 1 l of
purified water to the new blank. When all of the groups are finished- place the PCR tubes on the
cycler. Make sure that your PCR products are well-labeled so there is no confusion before the
next lab!
The program for the second PCR takes about 4 hours. This is because it employs a “touchdown”
method in which the annealing temperature is sequentially lowered by one degree from 52 to 47
degrees, running 2 cycles at each annealing temperature. Once 47 degrees is reached, the
program is run for 30 cycles. This increases the specificity and sensitivity of the primers. The
denaturing temperature (94 ºC) and the extension temperature (72 ºC) are the same as the first
PCR, and there is also a final extension at 72 ºC for 10 minutes.
The cycler is set to hold the samples at 4 º C after the program is finished. The TA will check
your PCR products on an agarose gel before the next lab to verify that the reaction worked. For
the second PCR, we expect a 200 bp product.
Clean up:
Save your first PCR product (well-labeled!) in the freezer, you will need this later in the semester
when we do cloning. 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:
8F:
5’ –AGAGTTTGATCCTGGCTCAG-3’
1
1492R:
5’-GGWTACCTTGTTACGACTT-3’
2
341F:
CGCCCGCCGCGCGCGGCGGGCGGGGCGGGGGCACGGGGGGCCTACGGGIGGCIGCA
3
533R:
5’-TIACCGIIICTICTGGCAC-3’
Lambda HindIII DNA Molecular Weight Standard (Ladder):
1.0% agarose
0.5µg/lane,
8cm length gel,
1X TAE, 17V/cm
*Note, under optimal conditions, an additional two bands at 72 bp and 125 bp may be seen.
1
W indicates an A or a T
This primer contains a GC clamp at the forward end to help with resolution when running DGGE
3
I indicates inosine base- helps increase sensitivity of primer
2
Lab 3: 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 the ERC for this lab. Also, this lab is going to be another long one, so be
prepared (I promise next week will be shorter :=)).
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 four teams will run their samples together on
one gel. Each gel can hold 16 samples, and each of the four groups has 4 samples. Also,
because about 2.25 hours are required to run the gel, the TA will prepare the gel in advance so
that we can load it and start it at the beginning of the lab. Then, we will prepare a gel together
while the first one is running so that you can see how it is made (like a cooking show!).
Procedure:
Running the Gel:
Washing the Wells:
When you arrive, the TA will already have prepared the gel and it 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 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 the heater and the pump
and remove the top portion of the electrophoresis unit 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.) positive control 2.) first
sample 3.) second sample 4.) negative control. You will use extra small-bore pipet tips which
allow you to reach the bottom of the well with the sample. Make sure when you take the sample
into the pipet 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 replace the top of the
Electrophoresis unit and turn on the heater and the voltage to 300 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 2
hours and 15 minutes at 300 volts.
Preparing the Gel:
Setting up the Glass Plates:
While the gel is running, you will learn how to prepare a gel. First the TA will demonstrate how
to prepare the glass plates for pouring the gel: First they are coated with SigmaCote 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:
100% denaturing solution
0% denaturing solution
Blue dye
H
13.2 ml
10.8 ml
425l
L
4.8 ml
19.2 ml
---------
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 gelonce 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 24 microliters of APS and 24 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 threeway 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.
After an hour- 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.
Preparing the Gel Stain:
Instead of Ethidium bromide we will use SybrGold nucleic acid stain. This stain is much more
sensitive than 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 (20 ul in 200 ml)
in 1X TAE buffer. We will keep this away from light until ready for use (it is photosensitive).
Staining the Gel:
Once the gel is finished, we will remove it from the unit, cut off the wells, notch the top right
corner (for orientation), and place it in the gel stain. We will then place it on an orbital shaker
(covered) for 15 minutes. While it is 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.
Imaging the Gel:
We will carefully transfer the 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. 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.
PM
b
Fig. 1: Image of DGGE gel comparing
microbial communities present in acid
mine drainage remediating
communities.
Lab 3: “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 randomly into a plasmid vector. 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. This
process is called transformation. Once the cells are transformed with a plasmid containing a
random 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.
Procedure:
GeneClean (Q Biogene) protocol for DNA purification:
First you must clean your PCR products. This removes any unused primers and dNTPs which
may interfere with the ligation. The geneclean kit works by binding the DNA to silica particles
(Glassmilk™), which allows you to “wash” the DNA with an ethanol-based solution (New
Wash™) while it is bound. Once the DNA is washed, it is eluted with purified water (GC
Elution Solution). This procedure should take about 10 minutes.
1.) Add 400 microliters of Glassmilk to a catch tube containing a spin filter (one for each
DNA extraction). Make sure to shake the glassmilk before adding to suspend the silica
particles.
2.) Add your DNA extract to the glassmilk and bind at room temperature for 5 minutes,
inverting tube every minute to mix.
3.) Centrifuge tubes at <14,000 g (Our centrifuge max is 13,300 g). Empty the flow-through
into a waste receptacle.
4.) Add 500 microliters of New Wash to the filter and centrifuge again.
5.) Discard flow-through and centrifuge the empty tube for 2 minutes to dry the glassmilk.
6.) Transfer the filter to a new catch tube and add 15 microliters of elution solution (purified
water).
7.) Centrifuge, discard spin filter. Your DNA is now purified! (Note- You could elute the
spin filter a second time for a 10-15% increase in DNA recovery (by mass)). Label this
final tube properly and proceed to the ligation step.
Ligation:
You will now insert your Genecleaned PCR products into plasmid vectors, which will then be
inserted into E.coli cells for cloning.
1.) Take 4 microliters of your genecleaned PCR product into a 200 microliter microtube.
2.) Add 1 microliter of salt solution and 1 microliter of 4-TOPO vector.
3.) Incubate the ligation reaction at room temperature for 15 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 degrees C and must thaw on ice
to avoid shock.
1.) Thaw the cells on ice just prior to use.
2.) Take 4 microliters 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 C and place back on ice for 5 minutes. During this
incubation, warm the SOC medium to room temperature.
5.) Gently add 50 microliters of the SOC medium to the competent cells. Incubate the cells for
15 minutes at 37 degrees C.
6.) Plate 100 microliters of the competent cells onto a prewarmed (at 37 C) Petri dish containing
LB agar and ampicillin. 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 ampicillin antibiotic will ensure that only cells which have taken up the 4-TOPO vector,
which contains an ampicillin 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.
Fig. 1: Map of 4-TOPO Cloning Vector
Lab 5: PCR of Clones and DGGE bands for sequencing
Overview:
We will now retrieve the inserts of your clones for DNA sequencing and identification using
primers specific to the plasmid vector. We 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.
At the same time- you will do PCR on the DGGE bands which you cut, using the water that they
have been soaking in as template. These are universal primers- so contamination is a concernbe extra cautions with these.
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.
Amount needed per 25 l reaction
Reagent
“Master Mix”: Amount needed
for N+1 Reactions (calculate)
16.15 l
2.5 l
5
l
0.5 l
0.25 l
0.25 l
0.35 l
Purified Water
10X buffer
5X buffer
dNTP (10 mM)
Primer M13F (20 M)
Primer M13R (20 M)
Taq Polymerase
Totals
25 l
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 which are medium to large in size, and which are well-separated. Do not
choose any tiny “pinpoint” colonies which are present, these usually grow in regions where the
antibiotic has degraded and do not contain the vector with the resistance gene.
You will then aliquot out 25 l of the Master Mix into 11 PCR tubes using pipet tips with a
white plug. Then, lightly dip a pipet tip into one of the colonies and dip 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. 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: 94ºC
10 minutes (this is longer than usual- both to break open the cells- and to denature the DNA),
followed by 30 cycles of 94ºC (15 seconds), 55ºC (15 seconds), 72ºC (15 seconds), followed by
2 minute extension at 72ºC. This program takes about 1.5 hours.
PCR of DGGE bands:
While the first PCR is running, set up a PCR for the DGGE bands. Each group cut out 5 bandsso prepare enough for 5 reactions plus the blank- or N+1=7, according to the following table.
Amount needed per 25 l reaction
Reagent
Purified Water
10X buffer
5X buffer
dNTP (10 mM)
Primer DGGEF
Primer DGGER
Taq Polymerase
“Master Mix”: Amount needed
for N+1 Reactions (calculate)
13.65l
2.5 l
5
l
0.5 l
1
l
1
l
0.35 l
Totals
24 l
Vortex the Master Mix and aliquot out 24 l into 6 PCR microtubes that have been labeled
according to the band number. Add 1 l of the water that the band has been soaking in into the
appropriate PCR tube. To the blank- do not add anything. Close the tubes and keep on ice.
Once the first PCR is finished- the TA will put the second set of PCR tubes on the thermocycler
and run the appropriate program (see lab 2 program for 341F 533R- these are modified versions
of the primers used in this lab and have the same program). In the next lab, you will purify these
products and prepare them for DNA sequencing.
Pick these
Fig. 1: Tips for choosing clones.
Avoid
these
Lab 6: Preparation for DNA Sequencing
Overview:
In this lab you will check your PCR products from both the clones and the DGGE bands from
Lab 5, gene clean them and determine the DNA concentration. This lab will be held at ERC so
that we can use the gel documentation system to determine DNA concentrations.
Checking PCR Products:
PCR products will be checked by agarose gel electrophoresis, as done in previous labs.
However, since we will be doing this lab at ERC, the TA will already have two agarose gels
prepared: one for all of the clone PCR products and one for all of the DGGE band PCR products.
Both gels will be made as before, except that SybrGreen will be used instead of Ethidium
Bromide to stain the gel. SybrGreen provides better resolution than ethidium bromide, which
will be useful in determining product concentrations. Also, it does not migrate towards the
cathode, as ethidium bromide does, which is useful when loading multiple rows of samples onto
the same gel as we will be doing.
Each group should take turns loading both their clone and DGGE band PCR products onto the
two gels. We will load the clone PCR products onto the larger gel since there are more of these,
and the DGGE bands products onto the smaller gel. Load 3 microliters of your product mixed
with 0.5 microliters of the dye. Also- a molecular weight size standard needs to be loaded in
each row.
We will run the gel at 150 volts for 30 minutes.
Documenting the Gel:
The gels will be documented using the UVP BioChemi Imaging system. This is the same high
resolution system which we used to document the DGGE gel. This system has a 16 bit camera
which supercooled to reduce noise. We will place the gels one at a time on the imager and
position so that it is in the middle of the field of view of the camera. Then we will make sure
that the filter is set on “SybrGreen” and capture an image of the gel.
This will be done for each gel. We will compare to the size standard in order to determine the
size of your insert. For the clones- we expect a PCR product of about 1700 bp (the size of the
16S gene plus part of the vector). For the DGGE bands, we expect a PCR product of around 250
bp. Also make sure that there is not any contamination in the blank- especially in the DGGE
band PCR, since this was done with “universal primers”. We will use the standard ladder in
order to verify the size of our PCR products.
Preparing for Sequencing:
You will now do a GeneClean on any PCR products which were the expected size. Do up to five
of the clones and all of the DGGE bands which worked. Follow the GeneClean procedure
described in Lab 4. You do not need to do an extra elution in the final step.
You will need to run the gene cleaned product on a gel one final time in order to estimate the
concentration. We will document the gel on the imager and use the software to estimate
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. 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 DGSeq, which has a melting temperature of 55 ºC and which we will
provide at 3.2 pmol/microliter.
The sequencing facility promises two day turnaround. Therefore we should have our sequences
returned in time for the next lab on sequence analysis.
Lab 7: Nuts and Bolts of DNA Sequence Analysis
Overview:
You will now analyze your sequences from both the DGGE bands and the clones and 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 receive a floppy disk containing their sequence files. The sequence files will be
named according to how you filled out the sheet- with an “.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. Also, the primer used for sequencing the
clones (T7) is specific to the cloning vector and is not susceptible to background contamination
present in your sample (The DGGE band primer used was a “universal” primer).
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. This 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-todate sequence information available on Planet Earth (that I am aware of). The advantage of
Blast is that it will allow you to check your sequence against this large 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. When this happens, you will see something like the 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 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.
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:
gi|30103112|gb|AY177357.2| Phenanthrene-degrading bacterium...
gi|23345134|gb|AY136080.1| Sphingomonas sp. KIN84 16S ribos...
gi|4868348|gb|AF131297.1|AF131297 Sphingomonas sp. JSS-54 1...
gi|12247763|gb|AF327069.1|AF327069 Sphingomonas sp. SA-3 16...
gi|456233|dbj|D13727.1|SPP16SRR6 Sphingomonas terrae gene f...
gi|30103121|gb|AY177366.2| Sphingomonas sp. 86 16S ribosoma...
gi|30060219|gb|AY254693.1| Uncultured alpha proteobacterium...
gi|40240919|emb|AJ619081.1| uncultured alpha proteobacteriu...
gi|19699044|gb|AY081981.1| Uncultured bacterium clone KRA30...
>gi|30103112|gb|AY177357.2|
ribosomal RNA gene,
partial sequence
Length = 1357
(bits) Value
337
337
337
337
337
329
329
329
329
7e-90
7e-90
7e-90
7e-90
7e-90
2e-87
2e-87
2e-87
2e-87
Phenanthrene-degrading bacterium M20 16S
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 common 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|
RNA gene, partial
sequence
Length = 583
Uncultured bacterium clone OSS-41 16S ribosomal
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:
One more important bit of sequence editing: For the clones, you will need to cut out the parts of
the sequence which belong to the cloning vector, and not to your sequence, and for the DGGE
sequences you may need to cut off parts of the primers. To do this, just look at the Blast
alignment and figure out where it starts and stops on the alignment. Export your corrected
chromatogram, open the file in Word, and cut off these parts where it starts and stops (This is
where it is helpful to use the Word “edit/find” function). Once you have your sequence editedsave 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 will 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/html/ and click on “Enter the Preview Site”. Now click on
“Sequence Match” and copy/paste your corrected sequence into the window.
Conundrum: In Blast, it was not necessary for you to know if your sequence was “forward” or
“backward”- Blast is smart and can figure that out. In RDP, however, you must know this. The
DGGE sequences will all be in the forward direction because they were sequenced with the
forward primer. The clones, however, are randomly inserted forwards or backwards into the
cloning vector. In order to figure out what direction your DNA was inserted- go back to the
Blast alignment and compare the numbers of “Query” (your sequence) with “Subjct” (the
sequence in the database). Are the numbers increasing for both? If so, your sequence was
inserted in the forward direction. If not, then it is backwards. In this case you will need to check
the box which says “Use the input sequences Complemented”. Everything else in RDP you can
leave as default, and finally click “Submit Sequences”.
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 the corresponding
RDP tree for the Blast alignment in Fig. 3.
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. For the clones- you have longer sequence
data, so you may have better luck with these.
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.
Band Name
Blast ID
2ea1-5
Alpha Proteobacteria,
Sphingomonas
(phenanthrene
degrading)
%
# bp
Ribosomal Database ID
match match
99%
170
Alpha Proteobacteria,
bp
Sphingopyxis/Sphingomonas
genus
Comments
Excellent
Sequence
Data
SEQUENCE_MATCH version 2.7
written by Niels Larsen.
Bacteria(domain)
Proteobacteria(phylum)
Alphaproteobacteria(class)
Sphingomonadales(order)
Sphingomonadaceae(family)
Sphingomonas(genus)
S000011084
S000017176
S000017317
S000018731
S000022926
S000123650
S000135888
S000143882
S000145112
0.945
0.914
0.908
0.945
0.914
0.933
0.945
0.945
0.908
1350
1318
1352
1350
1318
1196
1273
1322
1367
Sphingomonas sp. JSS-28; JSS-28; KCTC 2883; AF031240
uncultured soil bacterium; 749-2; AF423291
Sphingomonas sp. BF14; BF14 (bright yellow group 1 colony type) = DSM 9257; Z23157
Sphingomonas koreensis (T); JSS-26; AF131296
uncultured soil bacterium; 845-2; AF423296
uncultured alpha proteobacterium; APe4_19; AB074601
uncultured bacterium; C-CF-15; AF443567
uncultured alpha proteobacterium; KCM-B-125; AJ581589
Sphingomonas aurantiaca; MA405; AJ429238
0.982
0.951
0.926
0.908
0.939
0.975
0.982
0.939
0.982
0.939
1359
1389
1325
1360
1317
1325
1360
1333
1282
1282
Sphingomonas sp. SA-3; AF327069
uncultured bacterium; KRA30+14; AY081981
Sphingomonas macrogoltabidus (T); IFO15033; D13723
Sphingomonas sp.; IFO 15917; AB033950
Sphingomonas macrogoltabidus; IFO 15033T; D84530
Sphingomonas terrae (T); IFO15098; D13727
Sphingomonas taejonensis (T); JSS-54; AF131297
Sphingomonas adhaesiva; IFO15099; D13722
phenanthrene-degrading bacterium M20; M20; AY177357
Sphingomonas sp. 86; 86; AY177366
Sphingopyxis(genus)
S000004438
S000006063
S000012022
S000015501
S000015696
S000015765
S000020374
S000021987
S000146642
S000147004
Betaproteobacteria(class)
Burkholderiales(order)
Comamonadaceae(family)
unclassified
S000134527 0.945 1264 uncultured bacterium; C-FCF-16; AF443570
Fig. 7: Ribsomal Database Project classification of sequence presented in Figure 3.
Lab 8: Real-Time PCR
Overview:
Real-time PCR is a quantitative version of PCR which 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 meet at ERC for this lab
Real-time TaqMan PCR to Quantify Total Bacteria:
5’
BHQ
Cy-6
3’
1369F TM1389F 1492R
3’
5’
Fig. 1: Schematic of TaqMan PCR of Total Bacterial 16S rRNA gene
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. 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 for the SmartCycler are specialized to maximize the signal intensity (Fig. 2).
Setting up the Reactions:
Each group will set up duplicate reactions for each DNA extraction. The Smart Cycler
can process 16 tubes at the same time. We will therefore run 14 total samples (Group 4 has 1
extraction) and two of the blanks for the first run (the last two blanks will be run in a separate
run). 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
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
Reagent
Totals
Purified Water
10X buffer
5X buffer
dNTP (10 mM)
Primer 1369F
Primer 1492R
TaqMan Probe
Taq Polymerase
Mg 2+
Amount needed per 2524
ll
reaction
“Master Mix”: Amount
needed for N+1 Reactions
(calculate)
12.9 l
2.5 l
5
l
0.5
l
0.25
l
0.25
l
0.75
l
0.35 l
1.5
l
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
95 ºC
53 ºC
72 ºC
Repeat step 2-4 for 50 cycles
Smart Cycler cannot be set a
temperature lower than 45 ºC
2 minutes
15 seconds
60 seconds
20 seconds
Initial denaturing step
denaturing
Primer annealing
extension
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:
Y= -0.28X + 9.902
Y—Log [DNA]
X—Ct
Compare the value that you obtain with the original concentration of DNA that you had
calculated based on absorbance at 260 nm.