AFLP protocol

Last revised Sept 2000

            This protocol, currently used in the Wolf lab, has been adapted from several sources: The AFLP plant mapping guide from applied Biosystems (http://docs.appliedbiosystems.com/pebiodocs/00100509.pdf), the AFLP protocols of Laboratoire de "Biologie des Populations d'altitude" (Grenoble, France), and the KeyGene protocols by Matt Gitzendanner (Washington State University).

Restriction and Ligation reactions:

            The first step in preparing your samples for AFLP analysis is to cut the genomic DNA with two restriction enzymes (typically EcoRI and MseI). You will then ligate the adapters to the overhanging sticky ends produced by the restriction enzymes. There are two protocols described below; one protocol calls for the restriction and ligation reactions to occur in the same tube at the same time, and the other protocol requires that these two steps be done separately.

            Note: the restriction enzyme Tru9I is an isoschizimer (same recognition sequence and cut pattern) of MseI and is considerably cheaper. We have found that substituting Tru9I for MseI does not effect the results. Thus while the protocol still references the commonly used MseI enzyme we typically use Tru9I for our studies.  

AFLP oligonucleotide information:

PRIMERS: We've historically just used A's as the +1 nucleotides for 1+1

reactions. For 3+3 reactions, we generally use "A" plus some combination of

nucleotides that gives us a total of 3 G's or C's on both primers.

Getting the Adapters ready: (Note, this only needs to be done the first time that you mix the adapter pairs.)

            For each enzyme used, there is an adapter pair that will be ligated to the sticky ends. The adapters come as single strands, so the two strands of each adapter must be annealed to each other before they can be used.

1.   For EcoRI Adapter pair (final concentration of 5µM):

Mix:      25 µl of EcoRI Forward Adapter (@ 100µM)

             25 µl of EcoRI Reverse Adapter (@ 100µM)

           450 µl of TE                                                

           500 µl total

For MseI Adapter pair (final concentration of 50µM):

Mix:    250 µl of MseI Forward Adapter (@ 100µM)

           250 µl of MseI Reverse Adapter (@ 100µM)

           500 µl total

2. After mixing the adapters, heat at 95 ˚C for 5 min to denature. Then allow to cool slowly in a Styrofoam box to renature completely.

Setting up the reactions:

          For the restriction and ligation reactions, two master mixes are made up: one containing the enzymes, and one containing the adapters.  Use the reaction set-up sheets at the end of this protocol to set up the reactions. Here are some general comments regarding the restriction/ligation reactions:

1. Keep everything on ice as much as possible, to keep the enzymes from working before you want them to, minimize evaporation, and keep the adapters renatured.

2. Make sure you completely thaw and mix all buffers, especially the T4 DNA ligase buffer. Because the T4 ligase buffer has ATP, which is rather unstable, the stock tube of this reagent should be aliquoted it into small tubes to reduce freeze/thaw cycles.

3. A note on unit definitions of T4 ligase: there are actually two unit definitions for T4 ligase: Weiss units and cohesive-end ligation units. These are very different! In fact, 1 cohesive-end ligation unit (definition used by New England Biolabs) is only 0.015 Weiss units (definition used by just about everyone else). The thing is that the definition of a Weiss units is such that 0.01 Weiss units will ligate 1 µg of DNA in 20 min, so that’s about all you need in your reactions. In general, 1 cohesive-end ligation unit or 0.015 Weiss units per reaction is good.

3. The T4 Ligase is really concentrated, so you don’t need much and it is hard to pipette accurately. Don’t worry about this, as long as you get some in I think you’re fine.

4. Since the reaction volumes are small, and any evaporation could cause EcoRI to cut randomly (EcoRI* activity), in addition to cutting at specific sites, set up the reaction in 200 µl tubes and use a thermocycler with a hot top, or add  mineral for a conventional thermocycler. 

5. For the protocol that combines the restriction and ligation procedures, any fragment-fragment re-ligations that are made will be recut by the restriction enzymes. The adapter sequence is such that the restriction site is not regenerated following fragment-adapter ligation.

Below are two recipes for restriction-ligation; the first is the two-step procedure that we will follow in class (using the AFLP core kit from Life technologies).  This is followed by the one-step that we will follow subsequenctly when we buy the reagents separately.

I) Kit recipe.  First mix all the ingredients (minus DNA) enough for all tubes plus three extra (pipetting error).  Aliquot the mix, then add the DNA last, changing tips to avoid contamination across reactions.

Step 1. Digestion (per reaction tube):                                   

• 2.5 µl reaction buffer

• 1µl  EcoR1/MseI mix

• 50ng template DNA

• Distilled water to a total volume of 12.5µl

Incubate 2 hours @ 37 C

Then 15 min at 70 C to inactivate the enzymes

Step 2. Ligation, Add:   12µl adaptor/ligation solution

• 0.5µl T4 DNA ligase

• For a total now of 25µl per tube.

• Incubate @ 18-22 C ( room temp) for a further 2 hours, then store @ 4C

• Dilute 10-fold with distilled water

  II) Normal protocol (once we exhaust the kit):

      In a 0.2µl tube place the following (you can make a mastermix and aliquot later into the ligation tubes):

Enzyme master mix:

• T4 buffer 10X              0.1µl

• NaCl 0.5M                  0.1µl

• BSA @ 1mg/ml            0.05µl

• MseI: 1 unit                  0.1µl (usually – but check concentration of stock)

• EcoRI: 5 units               0.12µl (usually – but check concentration of stock)

• T4 ligase: 1 unit             0.2µl

• Distilled water               0.33µl

• Total per tube:              1µl

Then prepare ligation tubes (all except template DNA can be made as a master mix)

• T4 buffer 10X              1µl

• NaCl 0.5M                  1µl

• BSA @ 1mg/ml            0.5µl

• Enzyme master mix       1µl

• Mse adapter                 1µl

• EcoR1 adapter             1µl

• Total:                            5.5µl

• Template DNA             5.5µl (at about 10ng/µl)

  Mix well, centrifuge briefly to spin down droplets, and incubate at 37 C for 2 hours.  Store at 4C

  Add 90µl TE_0.1 to each tube (Note that the EDTA in the TE is at 0.1M)

+1 PCR Reactions (Preselective PCR):

            The first PCR reaction uses primers that match the adapter sequence and have one additional “selective” base. This reduces the number of bands that will be amplified. Theoretically this selective base could be any base, we use an A on the EcoRI primer (EcoRI+1A) and a C on the MseI primer (MseI+1C).

            The T4 DNA ligase only ligates one of the strands of the adapter to the fragment. The other is held on by base-pair binding to the other adapter strand. Thus the first step of the +1 PCR reaction is a 72 ˚C hold that allows the Taq polymerase to ligate the other strand. If you perform a Hot-Start, or place the +1 reactions into a hot thermal-cycler, or omit the initial 72 ˚C hold, you will loose the second strand and the PCR reaction won’t work. Do not use AmpliTaq Gold for +1 reactions as the Taq will not be active in the initial 72 ˚C hold.

 Use the set-up sheet at the end of this protocol to set-up the +1 reactions.

It is imperative that all solutions are thawed completely and mixed well when setting up the reactions.

In a 0.2µl tube (make a large mix as usual):

    Use the following PCR parameters:

1. 72 ˚C 2min

2. 94 ˚C 30 sec

3. 56 ˚C 30 sec

4. 72 ˚C 2 min

5. Goto 2 29 more times

6. 60 ˚C 10 min

7. 25 or 4 ˚C hold

  This profile is on the PE GeneAmp 2400 in the Wolf lab under “aflp-presel-30”

            Run 10 µl of the +1 reaction on a 1.5% agarose gel. You should see a smear in the 100 to 1,000 bp range. Sometimes bands are visible through the smear.

            Add 100 µl TE_0.1to the remaining reaction.

+3 PCR Reactions (Selective PCR):  

            For the final PCR reaction, the +1 products are used as template, and the primers are the same sequence except that they have two additional selective bases. The EcoRI+3 primers are labeled with a fluorescent dye (6-FAM in the Wolf lab) that can be detected by the ABI 377.

            Initially, you should test all EcoRI+3/MseI+3 primer combinations individually to identify the ones that work best (give the most readable and numerous polymorphic bands). If there are several EcoRI+3 primers that work well with one MseI+3 primer, and they are labeled with different dyes, then you can test them in multiplex reactions (where multiple EcoRI+3 primers are added to a reaction with a single MseI+3 primer) to verify that they produce that same bands as when they are run individually. If multiplexing does not work, it may still be possible to run separate PCR reactions, combine the products and run the samples in a single lane. However, as less of each reaction is loaded, the signal strength is reduced.

            It is imperative that all solutions are thawed completely and mixed well when setting up the reactions. This is especially important with the labeled primers as they tend to settle out of solution faster than regular primers.

Use the following PCR parameters:

1. 94 ˚C 2 min

2. 94 ˚C 30 sec

3. 65 ˚C 30 sec reduce by 0.7 ˚C per cycle)

4. 72 ˚C 2 min

5. Goto step 2 12 more times

6. 94 ˚C 30 sec

7. 56 ˚C 30 sec

8. 72 ˚C 2 min

9. Goto step 6 23 more times

10. 72 ˚C 10 min

11. Hold at 4˚C

This profile is on Carol von Dohlen’s PE 9700 under user = wolf, profile = aflp-select

  Running a gel:

            Running an AFLP gel on the ABI is very similar to running an automated sequencing gel. The same 5% LongRanger gel is used and all setup and loading is basically the same.

          Loading dye:

For each tube (make master mix as usual):

1. 0.75 µl of deionized formamide

2. 0.15 µl of blue loading dye (in GeneScan-500 box)

3. 0.3 µl of GeneScan-500 ROX size standard

4. 1.5 µl selective reaction

   Centrifuge briefly to get everything on the bottom of the tube.

   Heat at 95 ˚C for 2 min.

   Cool rapidly on ice and keep on ice until and during loading.

   Loading and running a gel:

   When setting up the gel, use the GeneScan Run and GeneScan Sample for setting up the run and sample sheet. For the sample sheet, you need to fill in the information for each lane, and each dye color in each lane.

   Load 1.5 µl  of sample on the gel.

   Run for 3 hrs.

   Settings to note on run:  Use the virtual filter set F:

   Modules:

• Plate check F

• Prerun: GS PR 36F 2400

• Run: GS run 36F 2400

 1. Open GeneScan.

1.1. Open the gel file you wish to analyze.

1.2.  Select Gel > Adjust Gel Contrast (command-J)

1.2.1. Move the top sliders down to increase the intensity of the color

1.2.2. Move the bottom sliders up to reduce the threshold for the color.

1.3.  Select Gel > Track Lanes…

1.3.1.  Click the Auto Track Lanes button.

1.4. When the tracker is done, check that the tracking lanes follow the lanes on the gel.

1.4.1. If the lanes are off, drag the tracking lanes to line them up with the lanes on the gel.

1.5. Note the scan numbers that correspond to the start and end of the data.

1.5.1. Select Settings > Analysis Parameters …

1.5.2. In the upper left set the start and stop ranges that correspond to those scan lines.

1.6. Select Gel > Extract Lanes … (command-L) and click OK.

1.7. The program goes through each lane and extracts the data, making an individual sample file for that lane. The sample files are all placed into a new folder within the folder the your gel file was in. Once all samples are extracted, the files are added to a project. Projects are used to collect samples for analysis, and can contain samples from multiple gels.

1.8. Make a size standard for the samples from each gel:

1.8.1. Select File > New… (command-N) and click on the Size standard icon.

1.8.2. Select a file to use for the size standard. You should pick one that worked well, and I generally choose one from near the middle of the gel. Click OK in the window that pops up next.

1.8.3. A new window comes up with the red portion of the chromatogram for your sample. This is the size standard peaks. Use the table and graph on pages A-6 and A-8 of the GeneScan Users Manual to identify the size of the peaks.

1.8.3.1. Click on a peak to assign a size.

1.8.3.2. Leave extraneous peaks as size 0.

1.8.4. Select File > Save … (command-S) and save your size standard. Call it something to indicate which gel and which lane the standard was generated from.

1.9. Go back to the analysis window, and click and hold on the pull-down box next to the "Size Standard" column header, and select the file you just created. This will apply the size standard to all sample in the project.

1.10. Click on the four color boxes to select all colors for all samples. Then click the analyze box.

1.11. GeneScan will now go through each sample and do the size calling for all peaks based on the size standard you created earlier. When it is done a new window will open up…the results control.

1.11.1. Double check the accuracy of size calling by examining all of the red data. If some samples do not line up correctly, make a new size standard for those samples and re-analyze them.

1.12. Save your project.

2. Open Genotyper.

2.1. Select Edit > Set preferences…

2.1.1. Select one color to import and click OK.

2.2. Select File > Import > From GeneScan File (command-I).

2.2.1. Select the files you want to import either one-by-one or click the import all button.

2.3. Click on the Samples section of the Genotyper window. And Select Edit > Select All (command-A) to select all of your samples.

2.4. Select Analysis > Clear Category List

2.5. Now we want to make a category that will determine which peaks will be scored. Only those peaks with a good signal in at least one individual will be scored. I usually set the good signal level at 300 fluorescent units.

2.5.1. Select Category > Add Category… (command-L)

2.5.1.1. Name your category something like: All peaks over 300.

2.5.1.2. I only look at peaks over 70bp so type 70 into the low Size box.

2.5.1.3. Click on the box next to the "with (scaled) height of at least" option, and type 300 in the text box after that.

2.5.1.4. Click OK.

2.5.2. Select Analysis > Label peaks, and click OK to label the peaks with the size in base pairs. This will label all the peaks with a fluorescent intensity over 300 units.

2.5.3. Select Category > Make from Labels… (command-M)

2.5.3.1. Unmark the "skip overlapping categories" box.

2.5.3.2. Type the prefix you want for the category names (color, primer set…)

2.5.3.3. Select the dye color that is being analyzed.

2.5.3.4. Click OK

2.5.4. Now you have a bunch a categories that correspond to the peaks that are over 300 fluorescent units in at least one individual. Now we need to score all of the individuals for those categories. To do that we need to label all of the peaks in all individuals (not just those over 300 units).

2.5.4.1. Select Category > Add Category… (command-L)

2.5.4.1.1. Name your category something like: All peaks.

2.5.4.1.2. Unselect the peak height  box.

2.5.4.1.3. Click OK.

2.5.4.2. Click in the categories box and select Edit > Unmark (command-U) to unmark all of the categories (notice that marked categories have a • in front of the category name). Then double click on, the All peaks category to remark it.

2.5.4.3. Select Analysis > Label peaks to label all the peaks. Click OK.

2.5.4.4. Now remark all the categories--select Edit > Mark (command-M).

2.5.4.5. Unmark the two categories that you made by hand (All peaks and All peaks over 300)

2.5.5. Now you are ready to make a table of the results.

2.5.5.1. Select Table > Add rows to table…

2.5.5.2. Click on the "More Choices…" button.

2.5.5.3. In the "Contents per row" set of radio buttons, select "Sample"

2.5.5.4. Click on the "Unmark All" button.

2.5.5.5. In the table setup box, double click on "Sample info" to select it (Now has a •)

2.5.5.6. Then double click on the "Labels" to select it.

2.5.5.6.1. The default setting for the labels is for each to take 2 columns in the table, but we only need one, so… in the settings box on the right, change the "Number of columns:" to 1.

2.5.5.7. Click OK.

2.5.6. Select Table > Export to file… and save the table.

3. Open Excel

3.1. Select File > Open…, and select your table, which is a tab delimited text file. Excel will present you with a bunch of options for opening the file, simply hitting return and accepting the default for each will work fine.

3.2. Modify your table in any way you want (shading every other row, bolding the header info, etc.…)

3.3. Print this out.

4. Open GeneScan.

4.1. Open your project

4.2. Select Window > Results Control (command-2)

4.3. Go through your samples (generally about 6 at a time) and for every category (now every column on your table) double check that you think that the sample was scored correctly.

4.3.1. If you think that there is a peak that was missed, make a note on you table.

4.3.2. If you think that a region is too hard to interpret make a note.

4.3.3. If you think two categories should be combined because they are really the same peak make a note.

4.3.4. Basically make any adjustments to the data that you see fit.

5. Go back to Excel and make changes.

6. Now you have a table with the manually checked calls for each peak for each sample. Now you can manipulate that data around in order to get something that a population statistics program will handle.

7. You are DONE!!!

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