Dna Barcodes Lab

Title

Application of DNA Barcodes to Identify Various Plant Species

Abstract
In this experiment we applied barcodes to plants in order to identify what species they are classified under. We also compared the DNA sequences of different plant species using the ribulose-biphosphate carboxylase gene (rbcL). We took samples from a plant called Chard and performed PCR, DNA amplification and quantification and sequenced the DNA. During the experiment, we hypothesized that this year’s “nonspinach” is Tatsoi, however, our results proved otherwise upon completion of a BLAST (see Fig 9). The completion of a BLAST showed that “nonspinach” is actually spinach (Spinacia Oleracea) and our sample was indeed Chard (Beta vulgaris, Fig 10). This confirms
the ability of the BLAST technique to identify and distinguish plant species.
Introduction

This experiment used the innovative technique of DNA barcoding. Barcoding is the process of using “short genetic sequence from a standard part of the genome to identify species. It works similarly with the way a supermarket scanner distinguishes products using the black stripes of the Universal Product Code (UPC). Two items may look very similar to the untrained eye, but in both cases the barcodes are distinct” (Hebert, 2003). According to Hebert, the most effective barcode regions in plants are found in the chloroplast (matK and rbcL).
Our main goal here was to distinguish the various plant species from each other by “using” short orthologous DNA sequences (barcodes)” (Kress et.al, 2005). The differences in the DNA sequences allowed us to distinguish which species belong where and how they are related. The plant species identified were Spinach, Radicchio, Frisee, Endive, Red Lettuce, Green Lettuce Chard and a hypothesized Tatsoi (nonspinach). In this exercise, we expect to find the greatest similarity between the same plants, between members of the same species, between the species in one genus, between the genera of same family and between the same order.
Methods

In this experiment, each group was given a bag of salad. Our group (B1 and B2) chose Chard as our specific plant type; however there were only two leaves present in the bag. As a result, my partner and I (group B1) shared one leaf. However, we sampled different sides of the leaf, in order to differentiate gene sequences (see Fig 1). Afterwards, we each isolated our DNA samples. We began by obtaining an isolation tube containing dilution buffer and crushed our plant sample in it using a 100 µl micropipette tip. We then took this mixture and spun it down in a centrifuge for 10 seconds to pellet the coarse plant fragments. Once we obtained the isolated DNA, we created a positive control and B2 created the negative control. After this was done, all the tubes were placed in a thermocycler.
A week after, our samples were taken out of the thermocycler and the isolated samples were tested for amplification by electrophoresis. The wells of the agarose gel were loaded with our PCR sample and loading dye (see Table 1 for map). After the wells were loaded, we began electrophoresis, setting the voltage at 110V. We then ran the gel for one hour, around the time when the dye reached 3-4 cm. Then we transferred the gel into a staining tray and stained it with Ethidium Bromide for 15 minutes. After 15 minutes, the gel was observed under a UV light machine. A photograph was taken in order to analyze the results of DNA fragmentation.
After the DNA has been amplified, it was purified. This was done using spin columns so the samples can be diluted. We accomplished the purification by adding Buffer PB and then observed the color changes in our sample. Since our sample immediately turned yellow, we moved on to its centrifugation and added buffer PE to wash the salts away. Buffer PE is made of ethanol and is used to wash the salt away from the DNA. We then added Buffer EB and centrifuged the sample again. DNA was quantified afterwards by using a standard size in electrophoresis. We loaded the PCR samples into the wells of the agarose gel (see Fig 3). After the results were obtained, we interpreted them using the size standard ladder provided in the protocol (see Table 2).
The last part of this experiment was sequencing the DNA.
Dr. Raleigh had the samples sent out to a lab in MIT for sequencing, but we performed a simulation of the sequencing process using paper clips. We began by synthesizing complementary strands of paper clips based on the base-pairing rules and also the template strand provided. We made copies of the template strand by lining it up with a complementary sequence. We paired an unpaired base I in the template strand and paid attention to whether or not it was “deoxy” or “dideoxy.” We continued with this process until we have a) picked a nucleotide (paper clip) with its 3’ end taped or b) we have completed a complementary strand. We repeated the process until there were no free nucleotides left in the pool. We then sorted the chains by length (shortest to longest), leaving the template strand out. The data from the whole class was pooled and can be seen in Table 3.
A week later, Dr. Raleigh received the sequencing results and distributed the results to the corresponding group (see Figs 4-6). A comparison between the forward and reverse sequences was made and the sequence was trimmed so a “clean” sequence was obtained. A clean sequence refers to a sequence whose primer and ragged ends were detached. A family tree was then constructed in order to establish relationships between different species. Only one sample from our group was used since the rest did not have enough material, so the other groups’ results were utilized as
a substitute.
Results:
Fig 1| Chard leaf sample


Fig 2| DNA Amplification by Electrophoresis


Fig 3| Quantification of DNA by Electrophoresis. Lane 2 was originally supposed to have our size standard (L), but the well was accidentally pierced, so we loaded Lane 1 instead.
Fig 4 and 5| Electropherogram (attached)
Fig 6| Nucleotide Arrangement (attached)


Fig 7| DNA Trimmed Sequence for 2011 and 2012 Chard

Figure 8| Phylogenetic Family Tree of 2011 and 2012 sequences


Figure 9| BLAST of “Tatsoi”


Figure 10| BLAST of ChardTable 1| Gel Map and Predictions on Fragments
Table 2| Mass of DNA in band for each dilution (ng)

Number of Bases COLOR OF LAST 3’ PAPER CLIP
“Wells” Table A Table B Table C Table D Table E Table F
17
16 5 Blue 4 Blue 4 Blue 4 Blue 1 Blue
15 1 Green 1 Green
14 1 Red
13 1 Yellow
12 1 Yellow
11 1 Blue 1 Blue 1 Blue
10 1 Green
9 1 Red 1 Red 1 Red 1 Red
8 1 Yellow 2 Yellow 2 Yellow
7 1 Green 2 Green 1 Green (1Yellow)*
6 1 Blue 3 Blue 1 Blue 1 Blue
Table 3| “Gel diagram.” *represents mutation

Discussion
Based on Figure 2, we were able to obtain the estimated band size of our PCR products.
Figure 2 also indicated the success of the PCR amplification since we were able to construct the gel map found in Table 1. In Table 1, lanes 1, 3, 6, 9,12 and 14 were expected not to show any DNA since they were not loaded. We used different primers for the positive and negative controls for the sole of validity and accuracy of the 100bp ladder when comparing bands in the experimental lanes. Our positive control was designed to amplify 297 base pairs. We expected to find 1 fragment we did. On the other hand, we did not expect any fragments to show in the negative control, since it did not contain any DNA and we were successful. In the lanes containing experimental samples, we expected to find a single band for each lane indicating that only one DNA fragment was amplified. On the basis of our gel, we can conclude that we were successful in amplifying the DNA of Chard because all of our lanes came out as expected. The experimental samples appeared to be identical, which makes sense because we amplified the same rbcL gene sequence fragment, which suggests identical banding patterns. Based on our gel alone, we cannot tell plant samples apart, but merely if the correct DNA fragment was amplified. For this region of DNA however, we expect to see some subtle differences between other plant specimens since some plants have more than one primer-bonding region. The rbcL fragments amplified in our
experimental sample tubes were ~600 base pairs in size, about twice the size of the positive control.
We were able to determine the concentration of our amplicon using the data from FIgure 3. We estimated the band size of our DNA based on our size standard (100 bp L). IN reference to Table 2, our estimated size for sample 5d2 was 21 ng, 5d4 was 16 ng, 6d2 was 24 ng and 6d4 was 13.3 ng. We were able to determine the original concentration of each of the PCR amplicons by dividing the mass (ng) by the volume of DNA solution (μl) added to the well and multiplying it by the dilution factor. After comparing the final concentrations, we observed that the numbers were relatively close, which tells us that our estimation was accurate.
Table 3 allowed us to to determine a complete gene sequence, since there was a general agreement that the color of the paper clip for each specific “well” number will always be the same. Our complete gene sequence was Blue-Green-Yellow-Red-Green-Blue-Yellow-Yellow-Red-Green-Blue. The basis for a new strand synthesis begins on the first nucleotide attached to the primer, hence why our table ends at 6 and not 1. Similarly, we are only looking at the last nucleotide in each chain to determine the sequence. The complementary strands we have made varied in lengths. 16-nucleotide chains appeared more so than others. We probably would have been able to increase the length, if we decreased the proportion of ddnNTPs in the reaction mix. Although we obtained a complete sequence, we only produced single-stranded chains. We were unable to produce many double-stranded DNA molecules in the process as we did in PCR because there was only one primer.
Upon completion of a BLAST for “Tatsoi” (Figure 9), the most likely genus for this species is Spinacia Oleracea due to a 100% match. This disproves the hypothesis that this year’s “nonspinach” is Tatsoi. This year’s “nonspinach” is actually “spinach.” In addition, after doing a BLAST for Chard (Figure 10), our sample was found to be of the genus Beta vulgaris. This finding confirms the ability of the BLAST technique to identify plants and is therefore a reliable source.
Literature-cited
Hebert, P. (2003). What is dna barcoding?. Retrieved from http://www.barcodeoflife.org/content/about/what-dna-barcoding Kress, W. J., Wurdack, K. J., Zimmer, E., Weigt, L., Janzen, D., & , (2005). Use of dna barcodes to identify flowering plants. PNAS, 102(23), 8369-8374.
Raleigh, F. Blackboard Shell. Documents, Notes, Power Points, and Outlines. Spring 2012 Semester. Saint Peter’s College.
Raleigh, F. Laboratory. Spring 2012 Semester. Saint Peter’s College.