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Mimulus guttatus Lab report

We constructed a double digest restriction associated DNA study on Mimulus guttatus. The first step was collecting samples, which we did on two field trips which can be read about here and here . Next, we extracted DNA from the samples we collected as well as samples collected by Alec detailed here. Next, we double digested our DNA using two restriction enzymes detailed here. These enzymes cut up the genome into many pieces. Next, we ligated unique DNA barcodes onto each of our individuals. The next step was using PCR for two purposes: to add a second unique and to test if our library construction was successful. Our PCR was successful as evidenced by a photo taken by Professor Paul. After the test PCR, we did a larger reaction of 25 micrometers that is identical to the previous one. This is the last step we were able to do as a class. In a perfect world, we would do the following other steps. The next step would be size selection. Size selection selects DNA of specific sizes, specifically, we would target ~400-600 bp. Size selection can be done in three different ways, one is using an automated system called pippin prep of which we have one housed in the Suni lab #Suni. A second way is to use gel extraction. Or, finally, magnetic beads can be used to isolate DNA. After size selection, we would then normalize our DNA samples, meaning bringing all of our DNA samples to approximately the same concentration. Having equal concentrations makes equal numbers of DNA fragments more likely to be sequenced. The final step would be to combine all of our size selected normalized PCR products into one vessel. Then, we would run these samples on any alumina sequencer- our class would run it on our in-house iSeq 1000 (Wall-e). Sequencing would take approximately 16 hours and, if successful, would generate tens of millions of reads. These data would be run through a bioinformatics pipeline. Ultimately, we would align these sequence data with the published Mimulus guttatus genome and call SNPs. Finally, we would use these SNPs to infer population differentiation using a metric like FST and assess population genetic diversity looking at things like the number of alleles, allelic diversity, etc. Based on what I know about Mimulus guttatus, I might expect populations that are genetically divergent. #MolecularEcologyForever

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Lab entry 12

After double digesting and ligating an adapter to the DNA, we did a test PCR. We did this to test for successful library construction of the samples.

  1. First, we made a master mix for the RADSeq using the following:
Master Mix Rxn: 1 Rxns: 11
NEB One-Taq 2x Master Mix 8 uL 88 uL
Forward Primer (10mM) PCR1X .4 uL 4.4 uL
Reverse Primer (10mM) PCR26 .4 uL 4.4 uL
Pure H2O 6.2 uL 68.2 uL
Master Mix Total: 15 ul 165 ul
Library DNA Template 1 ul  
Total reaction volume 16 uL  

 

2. We ran PCR1 on BIORAD #1/2 using 5 uL of the total reaction volume and 2 uL of loading dye.

3. Then, we ran the the products of PCR1 for each sample on 1.5% agarose gel with a 100 bp ladder at 130 V for 40 minutes.

The test PCR had samples #17-24.

For the final PCR, we used the same steps with the following master mix:

Master Mix Rxn: 1 Rxns: 11
Phusion DNA Polymerase .31 uL 3.41 uL
5X Phusion HF buffer 6.25 uL 69 uL
Forward Primer (10mM) PCR1_X 1.56 uL 17.2 uL
Reverse Primer (10mM) PCR2_1 1.56 uL 17.2 uL
DNTPs 10 mM .63 uL 6.93 uL
DMSO .94 uL 10.3 uL
Pure H2O 10.75 uL 118.25 uL
Master Mix Total: 22 ul 242.04 ul
Library DNA Template 3 ul  
Total reaction volume 25 uL  

We used the same samples #17-24. However, for samples 21 and 22 we used PCR27 as the reverse primer instead. This was a mistake made because we ran out of our original master mix and used another groups.

 

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Lab entry 11

Our next step for this lab was to do DD-RADSeq. First, we double digested our DNA samples. To do this, we followed the following steps:

  1. Double digested 100-1000 ng of high quality genomic DNA with selected restriction enzymes, using a digestion buffer appropriate for both enzymes.
  2. Then, I placed 6 uL of each sample’s DNA in the well of a PCR tube, storing it on ice.
  3. Then we prepared the master mix using the following measurements to make 130% excess of master mix 1:
    Master Mix Rxn: 1 Rxns: 11
    CutSmart buffer 10x .9 uL 9.9 uL
    EcoRI-HF enzyme .28 uL 3.08 uL
    MSPI enzyme .12 uL 1.32 uL
    Pure H2O 1.7 uL 18.7 uL
    Master Mix Total: 3 ul 33 ul
  4. We mixed it well, centrifuged it and stored it on ice.
  5. Then, we added 3uL of MM1 to each DNA sample
  6. We sealed the samples, vortexed, centrifuged, and incubated them at 37 degrees Celsius for 8 hours.

My samples were labeled #3 (SCHO002-26) and #4 (DIRA006-11).

Then, we did an adapter ligation. We did this using the following steps:

  1. First, we thawed the working stock EcoRI and Mspl adapters previously made as follows:
    PAUL LAB ID Adapter name
    Eco_2 AACCA_EcoRI
    Eco_3 CGATC_ EcoRI
    Eco_4 TCGAT_ EcoRI
    Eco_5 TGCAT_ EcoRI
    Eco_6 CAACC_ EcoRI
    Eco_7 GGTTG_ EcoRI
    Eco_8 AAGGA_ EcoRI
    Eco_9 AGCTA_ EcoRI
    Eco_10 ACACA_ EcoRI
  2. We added 1 uL of the working stock EcoRI adapter directly to the digested DNA.
  3. Then, we made a second master mix, making 130% excess with the following measurements:
Master Mix Rxn: 1 Rxns: 11
CutSmart buffer 10x .4 uL 4.4 uL
ATP (10mM) 1.3 uL 14.3 uL
T4 Ligase .2 uL 2.2 uL
Pure H2O .1 uL 1.1 uL
Universal P2 Mspl adapter 1.0 uL 11.0
Master Mix Total: 3 ul 33 ul

4. We added 3 uL of MM2 to the digested DNA

5. Finally, we sealed, vortexes, centrifuged, and incubated this at 16 degrees Celsius for 6 hours.

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Lab 10 entry- PCR Reactions

We used our DNA template to run PCR reactions (20mL). To do this, we created a master mix with the following solutions and measurements:

Ingredients per rxn per 18rxns (mL)
ddH20 13.36 240.48
10x buffer 2.00 36.0
MgCl2 2.00 36.0
BSA 1.00 18.0
dNTPs 0.20 3.60
F-primer 0.20 3.60
R-primer 0.20 3.60
Taq 0.04 0.80
Template 1.00 n/a
Total 20.00 19.00/rxn

First, I labeled three tubes with tube names QS10-12 respectively. Then, I used a pipette to put 19mL of the master mix in each tube. I then put each DNA template into its respective tube, changing pipette tips. Below is a list of tube names and their respective specimen ID.

Tubes Specimen ID
QS1MM MM1
QS2MM MM2
QS3MM MM3
QS4OY OY01
QS5OY OY02
QS6OY OY03
QS7 EB
QS8 RDRK001
QS9 SHOR005
QS10 KRS1
QS11 KRS2
QS12 KRS3
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Lab 9 entry

Once my DNA was extracted, I prepped it for gel electrophoresis.

  1. First, we dotted ~2mL of blue tracking dye on a piece of parafilm.
  2. Then, I put ~3mL of each extracted DNA on top of a dot, changing tips every time.
  3. Finally, I put my pipette on 6mL and moved the solution from the parafilm into the wells of the gel.
  4. We ran the gel. 
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Lab 8 entry- Modified Alexander et al. tube protocol for DNA extraction

  1. First, I labeled 3 2.0 mL tubes with my sample codes (KRS1-3 respectively).
  2. I added 3 sterile 3.2-mm stainless steel beads to each tube.
  3. Then, I added a small amount of leaf tissue into each tube, cleaning the tweezers between tubes to avoid contamination.
  4. Professor Paul loaded the tubes within a tube rack into the modified reciprocating saw rack and mounted the rack to the saw. He then turned it on speed 3 for 40 seconds.
  5. After, I centrifuged the tubes for 15-20 seconds at a fast speed to bring the plant dust down.
  6. I added 440 mL of preheated grind buffer to each tube.
  7. I incubated the buffered grandame at 65 degree C for 10 minutes in a water bath, mixing the tubes by inversion every 3 min.
  8. I then added 130 mL 3M pH 4.7 potassium acetate, inverted the tubes several times, and incubated the tubes on ice for 5 min.
  9. I centrifuged the tubes at maximum force for 20 min.
  10. I labeled new 1.5 mL tubes with the sample IDs and transferred the supernatant to the sterile tubes, avoiding transferring precipitate.
  11. Then, I added ~600mL of binding buffer to each tube, inverting to mix the solution.
  12. I added 650 mL of the new mixture to Epoch spin column tubes and centrifuged for 10 min, discarding the flow-through in a beaker.
  13. I repeated step 12 with the remaining solution.
  14. I washed the DNA bound to the silica membrane by adding 500 mL of 70% EtOH to the column and centrifuging at 15,000 rpm for 8 min until all liquid has passed through the membrane. I discarded the flow-through.
  15. I repeated step 14.
  16. Then, I centrifuged the empty columns at 15,000 rpm for an additional 5 min to remove any residual ethanol.
  17. I discarded the collection tubes and placed the columns in sterile, labeled 1.5 mL microcentrifuge tubes.
  18. Finally, I added 100 mL preheated pure sterile water to each tube and let it stand for 5 min before centrifuging for 2 minutes at 15,000 rpm. The solution left in the tube is the DNA.
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Lab 7 entry- Phylogenetic inference

I created an alignment of 25 COI sequences from Actinopterygii and my fish DNA barcode sequences, including one Chondrichthyes sequence as an out-group. I did this by searching COI and Actinopterygii/Chondrichthyes in the NCBI nucleotide program in Geneious. I chose sequences of similar lengths to my fish DNA sequences. I edited the alignment by choosing ‘Allow editing’ so that the sequences began and ended at the same point. When looking at the first 20 columns of my alignment, there were nine polymorphisms.

Best model based on AIC.

Best model based on BIC.

 

Then, I downloaded jModelTest2 to determine the best model of molecular evolution for my sequences. In Geneious, I exported the alignment in Philip format (relaxed). Then, in jModelTest2, I opened the file by clicking ‘File’ and ‘Load DNA Alignment’. Then I clicked ‘Analysis ‘and ‘Compute likelihood scores’, keeping the default settings. I looked at two methods, Akaike Information Criterion (AIC) and Bayesian Information Criterion (BIC). I did this by clicking ‘Do AIC/BIC’ respectively under the analysis window, keeping the default settings, and clicking ‘Do AIC/BIC calculations’. The best model based on AIC was JC, which was the same as the best model based on BIC.

Next, using Bayesian inference in Geneious, I selected my alignment, right clicked and chose ‘Tree’, then ‘MrBayes’. I used JC69 for the ‘substitution model’ and equal for ‘rate variation’. I kept the ‘gamma categories’ selection at 4. The ‘outgroup’ was put on HM422916, the sandshark outgroup I found earlier. For the initial ‘chain length’, I set it to 110,00. The ‘burn-in length’ was 10,000. I used the default setting for all other parameters. After running this analysis, I opened the ‘posterior output’ line and clicked on the ‘parameter estimates’ and ‘trace’ tabs.

Trace of MrBayes chain length 110,000

Parameter estimates of MrBayes chain length 110,000.

 

 

 

 

 

 

 

Next, using maximum likelihood to infer a phylogenetic tree, I installed RAxML. I chose a similar evolutionary model to the MrBayes model and chose ‘rapid bootstrap with rapid hill climbing’. Once RAxML was done, I right-clicked the new line and chose ‘Tree’ and then ‘Consensus Tree Builder’. Then, I clicked ‘create consensus tree’ and ‘support threshold’ of 50%. The clades in the resulting tree did not match the ones from the Bayesian analysis.

RAxML bootstrapping tree.

Initial Bayesian tree with bootstrap proportions.

 

 

 

 

 

 

 

 

 

 

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Lab 6 Entry

Using Geneious, we were able to match our fish DNA to DNA in the database. To do this we first copied the reverse sequence from the ‘Fish barcode Reverse Reads’ folder and pasted it into the ‘Forward Reads’ folder. Then, we selected the forward and reverse sequences of the same ID and clicked the Align/Assemble tab, choosing ‘De novo assembly”. Using the default settings, a new file appeared containing the consensus sequence for that fish DNA. We edited the sequence to clean up any discrepancies and saved these changes. We then right-clicked this file and chose ‘Generate consensus sequence’. We used this file to BLAST our sequence by right-clicking the file and selecting ‘BLAST’, keeping the default settings. Another file appeared, allowing us to see a set of top matches for our sequence. These matches told us what species our fish DNA most closely resembles. We created a new folder for a fish barcode test alignment and selected the assembly consensus sequence and 5-10 hits from the BLAST search, pasting these into the new folder. We then selected all of these sequences in the new folder and right-clicked, choosing ‘Multiple Align’ and then ‘Muscle Alignment’ with default settings. A new file was generated, showing polymorphisms between our DNA and other DNA of the same species.

Unfortunately, only two of my DNA barcodes were successful: KRS1 and KRS4.

Sushi Samples
Number Unique ID code Restaurant Species Name DNA Barcode Species Name
1 KRS1 Tuna Thunnus Albacares (Yellowfin Tuna)
2 KRS2 Mackerel N/A
3 KRS3 Albacore N/A
4 KRS4 Salmon Salmon Salar (Atlantic Salmon)

The successful DNA barcodes matched with the species specified at the restaurant.

For KRS1, there were 11 polymorphic sites found in columns 31, 167, 228, 324, 332, 333, 344, 346, 371, and 387.

For KRS4, there were 10 polymorphic sites found in columns 69, 586, 590, 611, 612, 614, 625, 628, 631, and 632.

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Lab 5 Entry- Second Field Trip

On Tuesday, September 24, 2019, we traveled back to Mount Tamalpais, this time stopping near Muir Beach to collect and view samples of mimulus guttatus. We stopped at a natural spring to drink the water and observe the flowering mimulus guttatus. We then traveled to a shaded creek bed to find more mimulus guttatus samples. Alec collected these samples to include in our lab.

Mimulus guttatus near a natural mountain spring (pictured behind the flower).

Mimulus guttatus flowers.

Mimulus guttatus in a creek bed.

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Lab 4- Gel Electrophoresis/ PCR Clean-up

To begin the electrophoresis of PCR product:

  • First, we thawed our PCR tubes
  • We dotted 16 loading dye dots (~1 ul) on a sheet of parafilm
  • We pipetted 3 ul of each PCR product on its own dot
  • We then loaded all dots into the gel (setting the pipette to ~5ul)
  • We ran the gel at 130 volts for 30 min

My PCR products were placed in lanes 9-13, beginning with KRS1 and ending with the negative control, respectively.

Clean-up of PCR products for sequencing- ExoSAP

  • First, we labeled new 0.2 ul PCR tubes with each of the sample codes
  • We then made the ExoSAP Master Mix
  • Lastly, we placed the PCR tubes into a thermocycler

ExoSAP PCR Clean-Up Protocol 

Recipe to clean-up one PCR reaction of 7.5 uL

Master Mix:                                  Rxn: 1          Rxns: 18

H2O                                                10.59 uL            190.62 uL

10x buffer (Sap 10x)                      1.25 uL             22.5 uL

SAP                                                0.44 uL               7.92 uL

Exo                                                0.22 uL               3.96 uL

Master Mix Total                    12.5 uL              225 uL

PCR Product:

PCR                                                7.5 ul

Total Cleaned-up Volume   20.0 uL

Steps:

  • We determined the number of PCR clean-ups
  • We calculated the volume of Master mix needed
  • We put reagents on ice
  • We pipetted 7.5 uL of each PCR product into a clean, labeled 0.2 uL PCR tube
  • Then we made the ExoSap master mix, keeping the reagents on ice while it was made
  • We pipetted 12.5 uL into each PCR product tube
  • We placed the tubes in a thermocycler and start the EXOSAP program
  • After the program completed (~ 45 minutes), we placed the PCR tubes in a labeled tube rack and placed them in the freezer

    DNA barcode for our group, Queen Salmon.

PCR results after 16 hrs 5 min.