Geneious II

October 11, 2018

During this lab session, Geneious was used to infer phylogeny trees for the 25 species of Actinopterygii that were chosen from the NCBI nucleotide database in comparison to the three sequenced nucleotide samples from the previous labs. Putting these together in one folder, a multiple alignment of all 28 sequences was created. Then, for better comparison between sequences, nucleotides tides were deleted from the beginnings and ends of the sequences to ensure equivalency in segment lengths using the shortest sequence as a basis for deletion. Six sequences did not fully match the homologous nucleotide sites of the others, so it was presumed that the misalignment may have been due to the sequences being the reverse complements. However, even after reversing the sequences and once again comparing all of the strands together, this misalignment was not solved. So, the sequences that did not align were removed from the overall alignment (though a different six sequences that matched the others were added later for the long-run analysis), which resulted in a end total of 22 sequences within that multiple alignment.

The next step was to install the jModelTest2 program in order to choose the best model of molecular evolution. Once the program was installed, the multiple alignment was exported in a Relaxed Phylip format and opened in the jModelTest program. The jModelTest then ran AIC and BIC analyses to generate likelihood scores for 88 models of molecular evolution to find the best model for the particular data set. Results from AIC analysis during the initial run of the experiment portrayed the best model to be TPM2UF + I + G, which did not correspond to the best model determined by BIC. BIC analysis showed HKY + G to be the best model of molecular evolution.

Afterwards, several hypothetical phylogeny trees were constructed based on results from Bayesian inference, maximum likelihood, and PHYML. Using the parameters outlined by the jModelTest2 and Bayesian inference, a short-run analysis was performed. The resulting posterior output graph did not have much of a shape or bars in general.

A second, longer run of the Bayesian inference using the same data was performed. The graphs obtained for this longer-run analysis showed a better distribution in terms of the bar graph shape and number of values. The predicted tree outcomes were very similar in terms of clades, but varied slightly in regards to the length of branches for specific sister taxas.

A maximum likelihood inference of a phylogenetic tree was also generated using the RAxML plugin on Geneious. Following the parameters of the evolutionary models determined by jModelTest 2 and choosing ‘Rapid bootstrap with rapid hill climbing’ , 100 trees were generated within one document. A consensus tree was then built out of these bootstrapping trees. However, the tree obtained was significantly different compared to the first analysis run with Bayesian inference since not only were many of the clades different, but the predicted max likelihood tree showed a significant amount of polytomy.

Finally, the last method tested used PHYML, but with the HKY85 model of molecular evolution  and the final Bayesian tree. However, despite running PHYML multiple times, the plugin seemed to have difficulty running and stays at 0% for over 15 minutes. Therefore, I was unable to obtain conclusive data using this particular method of computing maximum likelihood.

Geneious I

October 3, 2018

The introduction to Geneious lab segment illustrated the methods of utilizing the Geneious program. After completing the installation process, several sequences from the fish samples amplified in the previous labs were tested to identify whether the labels from the restaurant matched the information presented by the genomic data. These were the steps followed to complete the exercises using Geneious:

  1. Retrieved forward and reverse reads from Canvas and installed the files into Geneious.
  2. Copied and pasted a corresponding reverse read into the same folder as the forward read (eg. YH01_fwd, and YH01_rvs in same place).
  3. Assembled both forward and reverse reads onto same document using “De novo assemble”. Document was then named “YH#_ASSEMBLY”.
  4. Deleted any unreadable end bases.
  5. Modified bases that were illegible by comparing forward and reverse reads (eg. deleted unreadable bases on reverse strand and replaced it with legible base shown at homologous nucleotide site on forward read).
  6. Saved modified version of consensus sequence.
  7. Generated consensus sequence, which created new document under name “YH#_ASSEMBLY consensus”.
  8. Used BLAST on consensus sequence to search NCBI database for similar sequences.
  9. New folder of sequence matches appeared on menu.
  10. Identified whether sequenced sample matched labels provided at restaurant.

To build an alignment, these steps were taken:

  1. A new folder, named Fish Barcode Test Alignment, was created to host the barcode alignment documents.
  2. YH01_ASSEMBLY consensus sequence was copied and pasted into the newly created folder.
  3. 5 BLAST results within the top 100 hits for YH01 consensus sequence were selected and copied.
  4. The selected matching sequences were pasted into the Fish Barcode Test Alignment folder.
  5. All documents inside of the new folder were selected and the ‘multiple align’ function was used.
  6. A new document consisting of the nucleotide alignment of the five selected results and YH01 consensus sequence was generated.
  7. Polymorphisms were identified.

Due to unfortunate circumstances, the only samples sequenced using Geneious were YH01, YH02, and ARA01 (named YH_ARA01). YH01 was the sample of Tuna as depicted by the figure above, whereas YH02 consisted of a sample of Escolar. YH_ARA01 was an assigned sample of Yellowfin Tuna, as a replacement for the other samples that were unable to be properly amplified. YH01 and YH_ARA01 matched the species being served, though YH01 was not labeled as specifically. However, both samples were sequence-referenced through BLAST and the highest match percentages indicated that the species was Thunnus albacares, which was Yellowfin Tuna. On the other hand, YH02, the sample of Escolar, revealed a mixture of Thunnus albacares, and two different bacteria species. YH02 was sequenced three times, and broken into consensus sequences of YH02A, YH02B, and YH02C. BLAST results for YH02A matched the sample with Thunnus albacares, despite the differences in the sample’s appearance (Escolar versus Tuna), which indicated a degree of contamination between samples YHo1 and YH02. In addition, sequence YH02B matched most closely – though the percentage was fairly low – to Pesudomonas ludensis, a bacterial species that causes spoiling of milk, cheese, meat, and fish. Similarly, YH02C yielded a high match percentage with Pesudomonas fragi, a species of bacteria generally responsible for daily spoilages. YH02B and YH02C showed that not only was there contamination between samples, but also preservation issues regarding the sample of Escolar obtained.

According to the alignment built with YH01_ASSEMBLY consensus sequence, there were approximately 16 polymorphic sites (nt 681, 666, 426, 417, 375, 303, 288, 285, 279, 33, 25, 21, 12, 9, 6, 3) among the five chosen BLAST results. The first ten polymorphic sites were at positions: 3, 6, 9, 12, 21, 25, 33, 279, 285, and 288.

 

Field Trip II to Mt. Tamalpais

The second field trip consisted of an adventure through the winding paths of Mount Tamalpais. After passing through the Alpine Dam following the hiking trail, we climbed down a steep hill to find an area where the red-flowering mimulus cardinalis could have inhabited. The mimulus cardinalis is a flowering plant that generally uses hummingbirds as pollinators to reproduce.

 

 

 

 

 

The habitat consisted of a small, rocky, valley-like clearing with a small stream running through the side. Although the environment seemed to be a difficult place to inhabit, if a population of mimulus cardinalis was able to properly establish itself within this area, there would be significantly less competition due to the harsher conditions like the rocky bedding and limited sunlight. However, since the mimulus cardinalis’s main pollinators are hummingbirds, this area is still fairly accessible and may encourage reproduction despite the environmental conditions. Unfortunately, there were no populations that were spotted around this area, though there was interesting display of what appeared to be a ladybug festival.

We then moved across the bridge towards the other side of Mt. Tamalpais to another destination that usually had a population of mimulus cardinalis. The habitat was a small indentation at the side of the mountain, which was very accessible by pollinators, and surrounded by damp soil with a fair amount of sunlight. However, since the area was so small, there is a high possibility of in-breeding within this population of mimulus cardinalis compared to populations that inhabit larger environments.  

Afterwards, in order to observe the population of lupinus aboreus with purple flowers, we ventured to the other side of the mountain facing Stinson Beach. Atop the cliff-side, we were able to spot many individuals of purple-flowering lupine. The environment these individuals faced was very different compared to the populations at Pescadero Beach. While the yellow-flowering lupinus aboreus at Pescadero Beach had sandy soil, lots of sunlight, and fairly strong winds, the lupine at Mt. Tamalpais seemed to have little to no sunlight due to the heavy fog covering that side of the mountain, dry, grainy soils, and cold winds. The individuals at Mt. Tamalpais resembled the size of lupine at the cliff of Pescadero Beach, and appeared slightly smaller than those within the marshy area. The flowers also seemed to grow close to the ground in comparison to those at Pescadero.

                                      

Field Trip I to Pescadero State Beach

 

The first field trip on September 19th took the class to Pescadero State Beach, a coastal area approximately an hour away from San Francisco. We observed populations of Lupinus aboreus, a species of lupin that grows mainly along the coast of California. The Lupinus aboreus plant resembles a bush, growing relatively close to the ground with green finger-like leaves. However, the flower color of the Lupinus aboreus varies depending on its location, though there were only yellow flowers seen during the trip.

 

 

 

 

 

 

 

The coastal population that grew right next to the ocean atop a small cliff did not display any flowering, and the size of each plant seemed to be slightly smaller than the population that was growing further inland several meters. These small variations may be due to the fairly harsher conditions that these individuals faced, such as fierce winds and sandy soils compared to the more “sheltered” environment for those residing inland.

Finger-like leaves of Lupinus aboreus

Lupinus aboreus in the marshy inland

 

 

 

 

 

 

 

After crossing through the beach and under the bridge, we reached the population that was growing in the marshy inland near the beach. The individuals of Lupinus aboreus in this area displayed not only yellow flowers, but also seed pods that resembled edamame. The conditions seemed less harsh than the population growing by the beach as there was little wind, plenty of sun, and fairly moist soil.

 

 

The Sushi Test (cont.)

Subsequent to amplifying the genomic DNA (gDNA) collected from the fish samples, another gel electrophoresis was run, following the same steps as before, to see if specific bands were present. Unfortunately, some of the samples did not present the proper bands necessary to move onto the next procedure for PCR clean-up.

Another attempt was made by following the previous week’s steps for preparing the master mix and amplifying DNA via PCR. On the other hand, this would have been the protocol performed after obtaining the desired PCR results to clean up the amplified DNA and send for commercial sequencing:

  1. Prepare an ExoSap master mix containing water (10.59 μl), Sap 10x (1.25 μl), SAP (0.44 μl), Exo (0.22 μl), making sure that the reagents remain iced during preparation. The portions for each component should correspond with the numbers of reactions to be performed plus an additional amount covering for pipetting mistakes.
  2. Obtain a bucket of ice.
  3. Indicate and label the new PCR tubes to be used, then pipette 7.5 μl of each PCR product into the tubes.
  4. Pipette 12.5 μl of the ExoSap master mix into the newly labeled PCR tubes.
  5. Put the tubes in a thermocycler and start the ExoSap program, which should complete after approximately 45 minutes.
  6. Place the PCR tubes into a tube rack and put them in a freezer.

 

The Sushi Test

The Sushi Test illustrated the use of molecular tools and methods to amplify, sequence, and ultimately identify each sample’s species through its genomic DNA (gDNA). As such, fish samples were procured, disassembled for DNA extraction, run through Agarose gel electrophoresis, PCR-amplified, and commercially sequenced to discover whether the identities of the fish species corresponded to the labels used.

Collection

During the collection phase of the experiment, five sushi samples from Japanese restaurant Fujiyama-ya were obtained on September 4th at approximately 7pm. Using the cryo tubes provided, a very small portion of each fish sample was broken apart and placed inside of their respectively labeled containers. The samples were refrigerated until the day of the experiment.

Extraction

After obtaining the sushi samples, the next step involved extracting the DNA from the animal tissues. To do this, a DNA extraction kit, Sigma REDExtract-N-Amp Tissue PCR Kit, was utilized. This kit included three solutions: Extraction solution (ES), Tissue Preparation solution (TPS), and Neutralizing solution (NS). Other necessary materials were p200 micropipettes and micropipette tips, 1.5-mL microcentrifuge tubes, scalpels, weighing paper, balance, heat block, vortex, ice, sharpies, gloves, and ethanol solution. The following procedure was performed:

  1. Each sample was given a unique ID code, which was recorded on the “Animal Tissue DNA Extraction” data sheet. 
  2. After putting on gloves, a sharpie was used to label five 1.5-mL locking lid microcentrifuge tubes with the unique ID codes of the five samples. The ID code was written on both the top and the side of the microcentrifuge tubes.
  3. The samples were brought to a separate table to be prepared. Using a scalpel, a small piece of the sample was cut and placed onto a weighing paper.
  4. After taring the balance with the weighing paper, the cut sample was transferred to weight it with a goal of 2-10mg.
  5. The scalpel was cleaned with ethanol solution after each sample, and a different area of the weighing paper was used to disassemble different samples.
  6. Repeat steps three through five with the other four samples, or estimate using the amount portrayed by the first sample.
  7. Set aside the prepared samples.
  8. Using a p200 micropipette and an unfiltered tip, 100-μL of ES was added to each of the labeled microcentrifuge tubes.
  9. 25-μL of TPS was then added to the microcentrifuge tubes using the same pipette but with a new unfiltered tip.
  10. To mix the solution of TPS and ES, the mixture was pipetted.  This was repeated for all microcentrifuge tubes.
  11. Using foreceps, each sample was added to the corresponding tube containing the extraction mixture.
  12. The tissue sample was mashed inside of the tube using an unfiltered tip, making sure to change tips after each tube.
  13. Set the sample aside to incubate for ten minutes at room temperature.
  14. Put samples on a heat block and incubate the tubes at 95°for three minutes. Use a timer to ensure that the incubation time is exact.
  15. Remove the samples from the heat block.
  16. Using a p200 pipette and filtered tips, add 100-μL of NS and mix by pressing tube down onto the vortex machine.
  17. Place samples in ice, and check that the labels are still legible.

 

Agarose Gel Electrophoresis

The gel provided in this procedure was 1% agarose gel with GelRed added. Specifically, the composition of the gel involved 0.5g of agarose, 50-mL of 1x TAE buffer, and 0.5-μL of diluted GelRed for every 50-mL gel. The ensuing steps outline the set up of the gel box and sample preparation for gel electrophoresis to determine whether genomic DNA was obtained after the initial extraction:

  1. The gel tray was placed into the gel box with the top of the gel facing the red electrical connector.
  2. 1x TAE buffer was poured into the gel box wells, covering the gel with a small amount of bugger (about several millimeters).
  3. A medium-sized piece of Parafilm was cut.
  4. Using a p10 micropipette, pipette five dots of 2-μL Loading Dye in a row onto the Parafilm.
  5. Using the same p10 micropipette but with a filtered tip, add 3-μL of the genomic DNA from the microcentrifuge tube to the dot of Loading Dye. Make sure to switch pipette tips for each sample, and that each sample occupies their own respective dot.
  6. The genomic DNA added to the Loading Dye was equivalent to 5-μL, so the pipette reading was changed to 5-μL, and used to pipette the dots into the wells on the agarose gel. Record and take note of the wells for the respective specimens loaded.
  7. The lid was placed on the gel box with the colors of electrodes matching, and the power supply was turned on.
  8. The machine was set at 145 volts and run for approximately 15-20 minutes.

After running the gel electrophoresis, the machine was turned off and unplugged. The lid was taken off of the apparatus to expose the wells and gel box. Gently removing the gel box from the apparatus, the gel was moved to the imaging machine to be examined under UV light. The resulting picture produced by the imaging machine indicated that a good amount of genomic DNA was obtained from the samples of fish.

          

 

PCR Amplification of CO1 

Subsequent to confirming the presence of genomic DNA, the Sigma REDExtract-N-Amp Kit contained the necessary materials for PCR amplification, such as Taq polymerase, deoxyribonucleotide triphosphates, forward and reverse primers, and buffers that work with the materials from the extraction step. Additional materials include: purified water, p200 micropipettes, filtered micropipette tips, and a master mix containing the ingredients for PCR. Dilution of the genomic DNA was necessary to increase the efficiency of PCR because PCR works optimally with a minute amount of DNA template, whereas the creation of a master mix, which includes the sum of all reagents necessary for multiple samples, lessens the mistakes that can be made when pipetting small amounts. The following protocol details the gDNA dilution and making of the master mix necessary for PCR amplification.

  1. To make a 10:1 dilution of the gDNA obtained from the fish samples, microcentrifuge tubes were first labeled “1:10” along with the ID code of the samples including initials. Ensure that both the top and the side of the tube was labeled with this information.
  2. 18-μL of purified water was added to the five microcentrifuge tubes using p200 micropipettes.
  3. 2-μL of each genomic DNA was then added to the corresponding tubes.
  4. The solutions were mixed by flicking the tubes.
  5. Following the recipe featured below, a master mix was made.
  6.  Five PCR tubes were labeled on the sides and tops with the identification information of the gDNA samples.
  7. 2-μL of the 1:10 diluted gDNA was added to the PCR tube, then 18-μL of the master mix was added to the same tube.
  8. Repeat step 7 with all five samples, changing filtered tips between different gDNAs.
  9. Place the PCR tubes on ice until they can be placed in the thermocycler.

 

Skip to toolbar