Purpose:

The purpose of today’s lab was to discuss plans for the information that will be included on our poster and poster design. The goal was to discuss and correct any errors or misconceptions we had about how to write an “Abstract” for our research and to complete and edit the rough draft of our poster.

Procedure:

1. Discuss each group member’s abstract draft and work on perfecting it for the final abstract and title.
2. Go over poster design with Dr. Adair and work with group members to complete the draft of the poster. Finalize design and layout and include any information that is necessary for the presentation of our research.

Data:

Conclusion:

In conclusion, this lab was extremely productive and helpful. We were able to completely correct and finalize our abstract and title and everyone was productive and efficient when working on each part of the poster design. I was able to shorten the Introduction and Conclusion sections to bullet points and included References for our background information. Overall, we worked well together and we were able to complete the draft of our poster. The next step in this process would be to have someone from our group attend open lab tomorrow to edit the draft and see what we can improve upon. After that, we will be putting the finishing touches on our poster and presenting it in lab next week!

Purpose:

The purpose of this lab was to analyze the existence of DNA in the eDNA, positive control, and negative control samples that were amplified using PCR by running each sample through gel electrophoresis. The presence of a band would determine that the DNA extraction procedure and PCR amplification was successful and that DNA was present and prepared for further analyzation to determine the biodiversity of a sample. In addition, this lab was used as a time to prepare for upcoming poster presentations. Guidelines for posters, critiques and discussion of posters, and the beginnings of poster planning, design, and information needed on posters was discussed among groups.

Procedure:

Gel electrophoresis:

1. Place the agarose gel into the gel electrophoresis apparatus, with the wells closest to the negatively charged side of the box.
2. Pour in the previously prepared and obtained 1x TAE buffer solution onto the gel, filling the box and covering the gel and the wells completely.
3.  Collect your positive control, negative control, and eDNA samples from the front of the room. Carefully micropipette 10 µl of each sample into separate wells in your gel.
4. Micropipette 5 µl of DNA ladder already mixed with loading dye into the well adjacent to your samples.
5. Connect the positive and negative charged wires to their appropriate places on the box and to the power supply.
6. Place the lid onto the apparatus and turn on the power supply to the wires. Run the gel electrophoresis for about 30 minutes at 100 V.
7. Remove the gel after 30 mins and analyze under UV light to examine the bands of DNA present in the gel.

Data:

*Our wells were labeled 1-7, running from the left side of the gel to the right side. Well 1 begins on the left with our DNA ladder. We shared our gel with group 8 to run gel electrophoresis at the same time. Group 8 used wells #2-4 and our group (7) used wells #5-7.

Well #:

1. DNA ladder
2. Group 8 Negative control
3. Group 8 Positive control
4. Group 8 eDNA
5. Group 7 Negative control
6. Group 7 Positive control
7. Group 7 eDNA

None of the samples from our group yielded any DNA bands, however group 8 was able to see very light bands of about 450 bp for both their positive control and eDNA samples. Most of the class were able to see results on their gel electrophoresis and it was determined that the Chelex DNA isolation and extraction method was the most successful of all the methods that we had tested this semester.

Conclusion:

The lab went smoothly and all of the groups were able to complete the procedure for today’s lab with ease. Gel electrophoresis was a skill that we had already practiced and mastered in an earlier lab and we were able to use these skills to complete the procedure smoothly and efficiently in today’s class period. In addition, we were able to have meaningful discussion about our poster regarding its design, layout, and what information would be included on the poster. We were able to decide on a layout and delegated different sections of information on the poster to different group members, so that we could use our time and resources efficiently, while at the same time building each other up and relying on each other’s knowledge. We know what is expected to be completed by next lab time and can now plan accordingly for our next poster workshop day. In addition, when we were able to view the results from our gels toward the end of lab, we saw that our group’s samples yielded no DNA bands or positive results from our gel electrophoresis. These negative results could possibly be due to contamination of DNA that interfered in the PCR amplification process or possible errors of dilution or completion of procedure. There are multiple reasons that could explain why we got negative results, however it is difficult, if not impossible, to pinpoint the exact cause. While we did not see results, a majority of the other groups in our class were able to see results in their gels. Therefore, the Chelex protocol was determined to be the most successful at extracting DNA. When we were finished with lab, our PCR tubes and gel were stored back at the front of the room with the rest of the class and were labeled with the same marking information as referenced in the last log post. The next step for our lab group will be to create a poster presentation that will best describe our research that we have conducted this past semester.

Purpose:

The purpose of this lab was to perform the modified Chelex procedure one more time to see if we can obtain more positive results from our PCR samples than before and determine that this protocol would be the most successful when trying to determine a standard protocol to be used for metabarcoding. Based on previous extraction and PCR results, we determined that the Chelex protocol was the most effective, which is why everyone will be performing their own Chelex DNA extraction procedure and combining the DNA that we extract as a group to perform PCR on so that we can yield the most possible DNA sequences after amplification to be later sequenced and analyzed.

Procedure:

5% Chelex Protocol:

1. Using a compound microscope, view a non-flooded plate ant attempt to locate and isolate ciliates. Pipette 1.5 mL of ciliate sample water from the non-flooded plate and add to a small centrifuge tube.
2. Centrifuge the soil sample tube for 5 minutes at 6000 x g. Discard the excess supernatant.
3. Transfer another 1.5 mL from the non-flooded plate soil sample and add it to the soil sample tube.
4. Repeat step 2. This allows for you to analyze about 3 mL worth of DNA found in the water of the soil sample.
5. Measure out 0.5 g Chelex and transfer it to a 15mL conical tube. Add D.I. water up to the 10 mL mark in the tube. This step was already previously prepared for each group.
6. Add 200 µl of the 5% Chelex to the soil tube. Vortex for 1 minute.
7. Add 15 micro liters of proteinase K to the tube.
8. Incubate the tube for 30 minutes in 56 degree celsius heat block.
9. Once complete, place the tube in a 100 degree celsius heat block for 8 minutes.
10. Vortex the tube for 1 minute.
11. Centrifuge the tube at 16,000 x g for 3 minutes.
12. Carefully transfer 100 µl of supernatant to a clean centrifuge tube. Combine your supernatant with the 100 µl of soil sample from both of your lab partners. There should be about 300 µl of DNA sample in your combined tube.
13. Label the tube with your group’s information and store properly. Clean up all materials involved appropriately.

Preparing the PCR primer tubes:

Make 3 tubes of a 25 µl solution, each with V4 primers for PCR:

1. Obtain 3 separate centrifuge tubes, each already pre-made with 12.5 µl of 2X Master mix and ethidium bromide dye.
2. For the positive and environmental DNA tubes, add 1 µl of DNA template. For the positive control, this DNA comes from the DNA extracted from the Paramecium stock culture. For the eDNA, this is DNA extracted with Chelex protocol from your group’s combined tube. For the negative control, you will not add any DNA.
3. For the negative control, add 11.25 µl of D.I. water. To the positive and environmental DNA tubes, add 10.25 µl of D.I. water.
4. To each tube, add 1.25 µl of 10 µM stock V4 primers..
5. Label and store tubes for PCR amplification.

Making 1.8% Agarose Gel for Gel Electrophoresis:

1. Make 100 mL of 1xTAE in 1L Erlenmeyer flask by adding 10 mL of 10xTAE and 90mL of D.I. water.
2. Measure out 0.63g of agarose on a balance and add to a small erlenmeyer flask.
3. Add 35 mL of the 1xTAE solution to the small erlenmeyer flask.
4. Cover the flask with weighing paper and loose-fitting cap.
5. Heat solution in the microwave until it is clear and almost boiling.
6. Place in a water bath to cool the flask for about 5-6 minutes.
7. Add 2 µl of ethidium bromide.
8. Put together gel electrophoresis apparatus.
9. Gently pour the agarose solution into the box, avoiding any bubbles or other contaminants in your solution.
10. Allow to sit and solidify for about 30 minutes. Cover gel with some of the 1xTAE stock so that it will not dry out. Remove the comb and store in the refrigerator for later use.

Data:

Based on the nanodrop reading, the concentration of DNA in our sample was 473.6 ng/µl.

Conclusion:

We were able to complete all parts of the procedure in today’s lab, even though it took a little bit longer than expected. Although class ran a little long, we were able to complete more in this lab than we ever had before, showing that we have improved our understanding and technique of the procedure significantly over time and we are now able to confidently extract DNA using a variety of procedures, set up PCR tubes, and make an agarose gel for gel electrophoresis. We will run the PCR this week so that our next step during the next lab would be to run gel electrophoresis on our samples to see if the Chelex protocol was more successful and if it yielded more positive results as a whole. This will be helpful in allowing us to finalize our protocol for metabarcoding and we can then send our samples off to be analyzed to determine the biodiversity present among the amplified sequences.

Our DNA collection tube was labeled KGI 21-7 chDNA 4-5-18. Our PCR tubes were labeled 21-7 and the positive control, negative control, and eDNA tube were each labeled with a +, -, and e, respectively. Our PCR tubes were taken up to the lab to be amplified and our DNA collection tube was stored in a blue rack along with the rest of the class’s tubes.

Purpose:

The purpose of this lab was to analyze the results of our PCR from our tubes that we produced in an earlier lab. We ran our positive, negative, and environmental DNA samples through gel electrophoresis to determine if DNA was present in our amplified samples to see if our DNA extraction methods were successful or to determine if we needed to try a new extraction procedure if ours was unable to produce results.

Procedure:

• Gel electrophoresis:

1. Prepare 300 mL of 1X buffer solution using 10x TAE stock solution and D.I. water. Do this by measuring out 270 mL of D.I. water in a large graduated cylinder and adding it to a 500 mL erlenmeyer flask. Pipette 30 mL of 10x TAE stock solution and add to the flask using a serological pipette.
2. Place the agarose gel into the gel electrophoresis apparatus, with the wells closest to the negatively charged side of the box.
3. Pour in the 1x buffer solution onto the gel, filling the box and covering the gel and the wells completely.
4. Have each group member practice loading a well with a pipette, using whatever leftover diluted buffer as your sample.
5.  Collect your positive and negative control samples from the front of the room. Carefully micropipette 10 µl of each sample into separate wells in your gel.
6. Micropipette 5 µl of DNA ladder already mixed with loading dye into the well adjacent to your samples.
7. Connect the positively and negatively charged wires to their appropriate place on the box and to the power supply.
8. Place the lid onto the apparatus and turn on the power supply to the wires. Run the gel electrophoresis for about 30 minutes at 100 V.
9. Remove the gel after 30 mins and analyze under UV light to examine the bands of DNA present in the gel.

Data:

***Our wells were labeled 1-6, running from the right side of the gel to the left side. Well 1 begins on the right with our DNA ladder.

Well #:

1. DNA ladder
2. Negative control (COX 1 primers)
3. Positive control (COX 1 primers)
4. Environmental DNA (COX 1 primers)
5. Negative control (V4 primers)
6. Positive control (V4 primers)
7. Environmental DNA (V4 primers)

Conclusion:

Our group’s gel electrophoresis only showed results for the DNA ladder and the positive control using COX 1 primers. The rest of our wells showed no results. This could either be because the COX 1 primers were better primers to use for our experiment than V4, our DNA extraction method was not consistent, or our PCR protocol was contaminated or had something that prevented the DNA from being amplified properly. We will have to try our DNA extraction procedure again, or perhaps try the Chelex protocol instead to see if we obtain any better results. There could be a number of reasons that our gel electrophoresis showed that we didn’t have DNA in any of our other wells, however we will have to go through our procedure again to test new changes or try different techniques to see if they will prove to be more successful for our protocol in the future. Our tubes were stored in the box with everyone else’s PCR samples at the front of the room and are labeled Group 7. Our gel was disposed of after we had finished photographing and analyzing it.

Purpose:

The purpose of this lab was to continue with our protocol and conduct PCR on our environmental soil DNA that we extracted in last week’s procedure. At the beginning of class we discussed the details of how metabarcoding works and why we are choosing to specifically target the Cox 1 and V4 regions on the 18 S RNA. These regions are largely conserved, yet they differ in some areas for different organisms, making them useful in determining the presence of different species’ DNA in a sample. We are testing the efficiency of the two extraction methods with two different sets of primers in order to amplify the DNA we were able to extract from our soil sample.

Procedure:

Preparing the PCR primer tubes:

1. Make 6 tubes of a 25 µl solution, 3 of which have Cox 1 primers and 3 of which have V4 primers for PCR.
2. Obtain 6 separate centrifuge tubes, each already pre-made with 12.5 µl of 2X Master mix.
3. For the positive and environmental DNA tubes, add 1 µl of DNA template. For the negative control, you will not add any DNA.
4. For the negative control, add 11.87 µl of D.I. water. To the positive and environmental DNA tubes, add 10.87 µl of D.I. water.
5. To tubes 1-3, add 0.63 µl of Cox 1 primers and to tubes 4-6, add 0.63ul of V4 primers from a 20µM stock primer of each.
6. Label and store tubes for PCR amplification.

Making Agarose Gel for Gel Electrophoresis:

1. Make 100 mL of 1xTAE in 1L Erlenmeyer flask by adding 10 mL of 10xTAE and 90mL of D.I. water.
2. Measure out 0.63g of agarose on a balance and add to a small erlenmeyer flask.
3. Add 35 mL of the 1xTAE solution to the small erlenmeyer flask.
4. Cover the flask with weighing paper and loose-fitting cap.
5. Heat solution in the microwave until it is clear and almost boiling.
6. Place in a water bath to cool the flask for about 5-6 minutes.
7. Add 2 µl of ethidium bromide.
8. Put together gel electrophoresis apparatus.
9. Gently pour the agarose solution into the box, avoiding any bubbles or other contaminants in your solution.
10. Allow to sit and solidify for about 30 minutes. Cover gel with some of the 1xTAE stock so that it will not dry out. Remove the comb and store in the refrigerator for later use.

Data:

It is important to note that in the tubes labeled 1-3 we used Cox 1 primers and in the tubes labeled 4-6 we used V4 primers.

Each of the tubes are labeled with a -, +, or ED on top of the tubes, depending on if they contain negative controls, positive controls, or environmental DNA, respectively. They are also labeled with the initials “LR” on top. Each of the tubes contain the following PCR reactions:

Tube #1: Negative Control
Tube #2: Positive Control
Tube #3: Environmental DNA
Tube #4: Negative Control
Tube# 5: Positive Control
Tube #6: Environmental DNA

These tubes were stored in a rack at the front of the room with the rest of the class. In addition, our 1.8% 35 mL of agarose gel that we prepared and poured was left out on our lab table to dry and harden into gel. It is labeled with “Group 7.”

After running our sample through a nanodrop test, our DNA concentration from the environmental DNA we extracted was determined to be 10.942 ng/µl.

Conclusion:

In conclusion, we were able to complete the procedures for setting up the tubes for PCR with primers and creating the agarose gel for gel electrophoresis next week in good time during the class. After running a nanodrop test on the DNA we were able to extract using the MoBio Powersoil procedure, we were able to see that our results were pure and that DNA was present in the samples, meaning our extraction procedure worked. This was encouraging and we hope to be able to amplify this DNA for further examination so that it can be analyzed and the biodiversity of our soil samples can be assessed. The next step for our procedure would be to conduct PCR amplification on the tubes that we set up and run gel electrophoresis on the DNA from the tubes that we are able to amplify with the PCR.

Purpose:

The purpose of this lab was to troubleshoot where our previous Ludox DNA isolation protocol and EZNA DNA extraction protocol may have contained error that caused us to fail to yield any results of DNA being present in our samples. At the beginning of class and on the questions that matter, we discussed a number of reasons why this could have been the case, including contamination of ludox or ethanol in the samples, inefficient PCR reagents, or human error in completing the procedures in transferring the samples and conducting the DNA extraction protocol. After we discussed this, the groups then proceeded to conduct either the MoBio Powersoil protocol or the Chelex protocol in order to start the process over with a fresh start to try and experiment again to see if we could fix whatever problem might have caused us to fail to see results. Each lab table completed one MoBio procedure and one Chelex procedure, so hopefully we will be able to compare our results and see which DNA extraction technique was more efficient at determining the biodiversity of a soil sample.

Procedure:

MoBio Powersoil Protocol:

1. Measure out 0.25 g of soil sample and add to a PowerBead tube. In another PowerBead tube, add 250 micro liters of Paramecium stock sample. This will be your positive control. Follow the rest of the procedure for both the control and the soil sample.
2. Vortex the mixture briefly.
3. Add 60 micro liters of C1 solution. Vortex briefly.
4. Secure PowerBead tube to the MO Bio vortex adapter tube holder. Vortex at maximum speed for 10 minutes.
5. Centrifuge tubes at 10,000 x g for 30 seconds at room temperature.
6. Transfer the supernatant of the solution to a clean 2 mL collection tube.
7. Add 250 micro liters of C2 solution to the tube. Vortex for 5 seconds. Incubate at 4 degrees celsius for five minutes.
8. Centrifuge at 10,000 x g for 1 minute.
9. Transfer up to 600 micro liters of the solution supernatant to a clean 2 mL collection tube.
10. Add 200 micro liters of C3 solution to the tube and vortex briefly. Incubate for five minutes at 4 degrees celsius.
11. Vortex the tube at 10,000 x g for one minute.
12. Transfer the supernatant to a clean 2 mL collection tube.
13. Add 1.200 mL of C4 solution to the supernatant. Vortex it for 5 seconds.
14. Load about 600 microliters of solution in a clean tube and centrifuge tube at 10,000 x g for one minute. Discard the flow through from this process and repeat again. Add 600 micro liters of supernatant to the spin filter and centrifuge again at 10,000 x g for one minute. Load the rest of the supernatant onto the spin filter and centrifuge it at 10,000 x g for one minute.
15. Add 500 micro liters of the C5 solution and centrifuge it at 10,000 x g for 30 seconds.
16. Discard the excess fluid.
17. Centrifuge again at 10,000 x g for one minute.
18. Place spin filter in a clean 2 mL collection tube. (Avoid splashing c5 solution)
19. Add 100 micro liters of solution C6 to the center of the filter.
20. Centrifuge at 10,000 x g for thirty seconds.
21. Discard of the filter and label your tube with your group’s information. Store appropriately. Clean up work space and discard of materials appropriately.

Chelex Protocol:

1. Pipet 300-500 micro liters of soil sample into a small centrifuge tube. Gather another tube for your positive control and add the same amount of the paramecium stock sample.
2. Label both tubes and record your observations about the sample.
3. Centrifuge the soil sample tube for 5 minutes at 6000 x g. Discard the excess supernatant.
4. Transfer another 1.5 mL from the non-flooded plate soil sample and add it to the soil sample tube.
5. Repeat step 3. Repeat this process 3 times.
6. Measure out 0.5 g Chelex and transfer it to a 15mL conical tube. Add D.I. water up to the 10 mL mark in the tube.
7. Add 200 micro liters of the 5% Chelex to the soil tube. Vortex for 1 minute.
8. Add 15 micro liters of proteinase K to the tube.
9. Incubate the tube for 30 minutes in 56 degree celsius heat block.
10. Once complete, place the tube in a 100 degree celsius heat block for 8 minutes.
11. Vortex the tube for 1 minute.
12. Centrifuge the tube at 16,000 x g for 3 minutes.
13. Carefully transfer the supernatant to a clean centrifuge tube.
14. Label the tube with your group’s information and store properly. Clean up all materials involved appropriately.

Data:

Our lab group completed the MoBio Powersoil DNA protocol. We used 0.3 grams of soil for our experiment, since our scale could not weigh out exactly 0.25 grams. At first when we were trying to separate the supernatant from the soil after we centrifuged, it was difficult to collect any supernatant from the sample because the soil had a sludgy texture to it and there wasn’t much liquid to collect. However, after we centrifuged it again at 10,000 x g we were able to collect more supernatant off of our sample, making it a more reliable and error-free experiment. We completed the rest of the procedure, labeled our tubes G7, and stored it in the freezer on the white rack.

Conclusion:

Although it is frustrating that we were not able to yield any positive results of DNA in our samples from our previous experiment, I am hopeful that these new protocols will yield positive results if we perform the procedures correctly and carefully. The instructions are clearly laid out and by this point in the semester we have mastered the basic laboratory techniques needed to complete the procedure, since we have gone through other protocols similar to these ones earlier. Our next step in our experiment is to proceed with the PCR of our samples.

Purpose:

The purpose of this lab was to continue working on our gel electrophoresis procedure from last week’s lab. Since we had already prepared the gel and our samples last week, all we had to complete this week was the actual process of running the samples through the gel and observing if any samples were present and what size they are by comparing it to the DNA ladder that we also ran. We discussed the different uses for agarose gel electrophoresis and reviewed that we would be using gel electrophoresis in order to determine the presence/size of amplicons from our PCR DNA sample.

Procedure:

1. Class was started by discussing the various uses of gel electrophoresis and stating our purpose of conducting gel electrophoresis on the positive and negative controls and our soil sample after it had been amplified by PCR.
2. We then began our gel electrophoresis:
   1. Prepare 300 mL of 1X buffer solution using 10x TAE stock solution and D.I. water. Do this by measuring out 270 mL of D.I. water in a large graduated cylinder and adding it to a 500 mL erlenmeyer flask. Pipette 30 mL of 10x TAE stock solution and add to the flask using a serological pipette.
   2. Prepare 30 µl of 1X loading dye. Do this by adding 5 µl of 6X loading dye and 25 µl of D.I. water to a small tube. Mix well.
   3. Place the agarose gel into the gel electrophoresis apparatus, with the wells closest to the negatively charged side of the box.
   4. Pour in the 1x buffer solution onto the gel, filling the box and covering the gel and the wells completely.
   5. Have each group member practice loading a well with a pipette, using whatever leftover diluted buffer as your sample.
   6.  Collect your positive and negative control samples from the front of the room. Add 5 µl of bromophenol blue dye to each of your samples. Mix well. Carefully micropipette 10 µl of each sample into separate wells in your gel.
   7. Micropipette 5 µl of DNA ladder already mixed with loading dye into the well adjacent to your samples.
   8. Connect the positively and negatively charged wires to their appropriate place on the box and to the power supply.
   9. Place the lid onto the apparatus and turn on the power supply to the wires. Run the gel electrophoresis for 30 minutes at 110 V.
   10. Remove the gel after 30 mins and analyze under UV light to examine the bands of DNA present in the gel.
3. Complete the QTM with you group.
4. Review your group members’ rough draft Introductions and critique on what could be improved. This will be reviewed in further labs.

Data:

While we ran our gel electrophoresis, we were able to work on some practice problems for dilutions and how to prepare the buffer that the gel would be sitting in and the current would be conducted through in gel electrophoresis. For example, when we were preparing our buffer solution, we needed to make a 300 mL 1x dilution buffer using 10x TAE stock solution and needed to determine how much D.I. water we would add to the TAE stock solution in order to achieve this solution. In order to solve the problem, you would use the equation C1V1 = C2V2. The stock solution and water needed to be in a ration of 1:9 and we calculated that we needed 30 mL of TAE stock solution and 270 mL water to create this dilution buffer for our gel electrophoresis. In addition, we discussed what future steps we would take after gel electrophoresis, should we get positive or negative results. If we got positive results, this meant that DNA is present in the sample and we would send the sample to a company to be counted and sequenced. If we got negative results and no bands were present, we would have to go back through our procedure to see where we might have gone wrong and make some changes in the future to see if we can get results.

We were not able to finish the gel electrophoresis in lab because we ran out of class time, so we left our materials out and still running on our table when we left. Dr. Adair completed our gel electrophoresis and got pictures of our gels, however none of our samples in our gel yielded any results. There could be a number of reasons for this, but ultimately we will just have to go back through our procedure again and brainstorm what might have caused this to occur.

     .  

Conclusion:

Although we didn’t get to complete our gel electrophoresis procedure ourselves, we were still able to get valuable practice in making dilutions and were able to complete most of the procedure. When our gels had finished running through gel electrophoresis and were illuminated by UV light, we were unable to see any positive results of DNA strands being present in the gel, even in our positive control. There are many possibilities of why this occurred and we will have to trace back our steps to figure out how we can improve our procedure. I would guess that something went wrong with our DNA extraction procedure that caused our samples to have no DNA in it. Some future steps that we will take in the following lab is going back through our procedure and thinking through and possibly changing some steps to see if we can fix whatever problem occurred and determine what might have caused this. We may have to try a different DNA extraction protocol entirely.

Our gel electrophoresis apparatus was left on our lab table so that the procedure could be completed and taken up to a different lab to get pictures of our gel. We returned our negative and positive control and our soil PCR DNA samples to the rack at the front of the room that they were placed in last week. They are labeled with a “7” on the lid and are located in slots A1, A2, and A3.

Purpose:

The purpose of this lab was to prepare both positive and negative control samples, as well as our soil DNA samples in tubes to be put through the PCR amplification process of denaturation, annealing, and elongation. In addition, we discussed and learned the details of the PCR amplification process to ensure that everyone in the class had a clear understanding of how we would be amplifying the DNA we found in our samples. We discussed the possibility of skewed results, due to our unreliable and questionable results of testing for DNA in our samples. It is possible that our cell layers were contaminated by other substances in the DNA extraction procedure and this is what is causing our results to be unreliable and unusual. Therefore, when we run PCR it is possible that other cells’ DNA, not just ciliates, will be amplified in the process as well. Lastly, an objective of this lab was to make an agarose gel that we would be using for gel electrophoresis on our amplified DNA samples in next week’s lab.

Procedure:

1. Discuss Learning Objectives of the lab and the details behind the PCR amplification process.
2. Discuss the results of analyzing our DNA concentrations from the samples that we were able to extract using the procedure from last week’s lab.
3. Begin preparation for the PCR procedure:
   1. Obtain 3 tubes already filled with 12.5 μl of 2x Master Mix. Label each of these tubes with a +,-, and S to indicate which tube will be the positive control, negative control, and soil DNA sample tube. The master mix consists of the appropriate buffer, DNA nucleotides, and Taq polymerase. Perform each of the following steps quickly and carefully, so as to limit the amount of DNA exposure and contamination throughout the experiment.
      1. For each of the tubes, add 1 μl of COX 1 forward and reverse primers using a micropipette.
      2. For the negative control tube: pipette 11.5 μl of deionized water into the solution. No DNA template will be added to this tube. Set aside once its preparation has been completed and it has a total of 25 μl.
      3. For the positive control: pipette 5 μl of the DNA template stock solution of Paramecium (concentration of 10 ng/mL) into the solution. In addition, pipette 6.5 μl of deionized water into the solution, to reach a total of 25 μl in the tube. Set aside for storage.
      4. For the soil DNA sample tube: pipette 5 μl of the extracted DNA from our soil samples into the solution (concentration unknown and unreliable.) In addition, pipette 6.5 μl of deionized water into the solution, to reach a total of 25 μl in the tube. Set aside for storage.
      5. Place the three tubes in the class rack at the front of the room. Record the location of your tubes in the rack and what you indicated each of them with. Our tubes are labeled with a “7” on the lid and are located in slots A1, A2, and A3 in the rack.
4. Begin making Agarose gel:
   1. Measure out 0.6 grams of Agarose. Add to 40 mL of T.A.E. Swirl the erlenmeyer flask with the solution to begin to mix. Cover the flask with weighing paper and a small plastic cap to prevent contamination.
   2. Microwave the flask apparatus for 1 minute and 20 seconds. Remove carefully, solution is boiling.
   3. Place flask into a water bath at 55 degrees Celcius for 5 minutes to allow it to cool.
   4. Add 2 μl of ethidium bromide to the flask, swirl the solution to mix.
   5. Once mixed, pour the solution into a gel rack to sit, cool, and harden into gel to be used for gel electrophoresis next week. Store away in proper place in the classroom and clean your lab area.

Data:

Once we completed our preparation for the PCR of our control tubes and soil sample tube, we began to make the agarose gel that we will be using for gel electrophoresis in next week’s lab. In order to create a 40 mL 1.5%  T.A.E. solution agarose gel, we measured out 0.6 grams of agarose and added it to 40 mL of T.A.E. and swirled to begin to mix the solution. We microwaved the solution for 1 minute and 20 seconds, then let the erlenmeyer flask of our solution cool in a water bath at 55 degrees Celcius for 5 minutes. Then, we added 2 μl of ethidium bromide to the flask, swirled the solution to mix, then poured the solution into the gel rack to sit, cool, and harden into gel to be used for gel electrophoresis next week.

Each of the tubes from our PCR procedure are labeled “7” on the top of the lid and are labeled “+”, “-“, and “S” on the sides of the tubes for the positive control, negative control, and soil DNA tube, respectively. The positive control tube, negative control tube, and soil DNA tube were each placed in slots A1, A2, and A3 in the class’s rack, respectively. My group’s gel rack is labeled “Group 7, 1106-21” and has each of our initials: LHR, SGS, and KGI written on a piece of tape that is placed on the side of our gel rack. The gel rack was stored along with the other gels the other groups had made and was let out to sit and harden into a gel-like jelly on top of our lab counter. We cleaned up the rest of our lab station and returned all of the materials we used to their proper place in the lab room. We cleaned out our erlenmeyer flask with bleach and hot water after pouring our agarose solution into the gel rack. The flask was placed to the side on the counter next to the sink to dry. Below is a picture of our gel solution just after we finished pouring it into the gel rack. 

Conclusion:

We were able to work quickly and efficiently in this lab and we were even able to finish the lab procedures before the lab class time was finished. We had some good discussion at the beginning of class about the specifics of PCR amplification and we were able to clear up any confusion or misconception that we had on that topic.The discussion time at the beginning of class was extremely helpful because it allowed me to clearly understand the purpose of forward and reverse primers and how this process could allow us to sequence several copies of strands of DNA, specifically targeting the v4 region of the 18s rRNA We were also able to make agarose gel to be used in gel electrophoresis in next week’s lab. This was useful because the next step in our experiment would be to run gel electrophoresis for each of our control and soil DNA samples that we are able to amplify once we complete the PCR process. This will allow us to visualize our DNA strands and estimate their size and length in nucleotides when we compare them to the DNA ladder we will also run. I am looking forward to completing this procedure in next week’s lab!

Purpose:

The purpose of this lab was to complete the rest of the Ludox procedure that we had edited and finalized and had already begun last lab, as well as to learn about and begin the EZNA DNA extraction kit protocol. We first went over the EZNA DNA extraction kit protocol in detail so that we would be very familiar with the process so that it would run smoothly, effectively, and efficiently. The main focus of this lab today was to begin moving on from the DNA isolation part of our metabarcoding process and progressing to the DNA extraction part of our experiment. We will be moving on from using our Ludox procedure and will proceed to take our samples from the Ludox procedure and extract the DNA from our samples using the EZNA extraction kit so that the DNA can be amplified and examined in future labs.

Procedure:

1. At the beginning of lab, the class discussed the EZNA DNA extraction kit protocol in detail. This ensured that all of the groups knew exactly what was expected of them during the lab so that we could perform the procedure effectively and efficiently, without diving into the procedure before we actually knew what was needed for the DNA extraction to occur. Students were able to ask questions and went over the details of each step, walking through the different materials and techniques they would be using during the procedure.
2. Student then began to complete the Ludox procedure from the previous lab, using the pelleted samples that were stored in the freezer from the previous week. Students completed DNA isolation using the Ludox cell layer by completing the final half of the procedure that was as follows:
   1. Spin both of the 2 mL tubes with Ludox cell layer in centrifuge for five minutes at 3000 g.
   2. Remove the excess supernatant until only the DNA pellet remains. Be careful not to disturb the pellet.
   3. Add 1 mL of 1x PBS. Resuspend the DNA pellet in the buffer in order to wash the cells.
   4. Spin both of the 2 mL tube using the centrifuge again for five minutes at 3000 g.
   5. Remove the excess supernatant until only the DNA pellet remains.
   6. Resuspend cells in 100 µl PBS in each tube. Combine the samples from both tubes into one tube so that you have a total of 200 µl of solution.
   7. Remove 20 µl of cell sample from 2 mL tube and pipette into a smaller holding tube. Add 20 µl of Iodine to the 20 µl of cells and mix well.
   8. Have each member of the group remove at least five 2 µl drops from the iodine-cell mixed sample. Place these drops on a concavity slide and view using a compound microscope. Observe and record the number of cells you are able to see in the drop.
3. Next, begin to go through the steps of the EZNA DNA kit extraction protocol that has been modified for ciliate samples. The following is a summarized procedure of the EZNA kit protocol:
   1. Before Starting: Set heat block to 70°C. Heat Elution Buffer to 70°C in the heat block. Chill PBS to 4°C by placing on ice.
   2. Prepare the cell suspension: Wash the cells with cold PBS by adding 200 μl PBS, resuspend the cells using the vortex or flicking, and then spin with the centrifuge to return to pellet form. Remove the supernatant.
   3. Resuspend cells in 200 μL of PBS.
   4. Add 25 μL OB Protease Solution. Vortex to mix the solution thoroughly.
   5. Lysis: Add 220 μL BL Buffer.
   6. Mix well with vortex before placing the tube on the heat block to incubate at 70°C for ten minutes. Briefly vortex the tube in the middle of incubation when about five minutes have passed.
   7. Binding: Centrifuge tube for a couple of seconds. Add 220 μL ethanol. Vortex to mix solution well.
   8. Insert a HiBind. DNA Mini Column into a 2 mL collection tube.
   9. Transfer the entire sample to the HiBind. DNA Mini Column, including any precipitates that may have formed.
   10. Centrifuge the tube at maximum speed for 1 minute.
   11. Discard the filtrate and reuse the collection tube.
   12. Wash and Dry: Add 500 μL HBC Buffer to the column.
   13. Centrifuge at maximum speed for 3o seconds.
   14. After the HBC Buffer wash, discard filtrate and collection tube.
   15. Insert the HiBind DNA Mini Column into a new 2 mL collection tube.
   16. Add 700 μL DNA Wash Buffer.
   17. Centrifuge at maximum speed for 30 seconds.
   18. Discard filtrate and reuse the collection tube.
   19. Repeat steps 16-18 to perform a second DNA Wash Buffer.
   20. Centrifuge the empty HiBind DNA Mini Column at maximum speed for two minutes to ensure that it is completely dry.
   21. Elute: Transfer the HiBind DNA Mini Column into a 1.5 mL microcentrifuge tube. Label the tube with your lab group’s identifying information.
   22. Add 100μL Elution Buffer heated to 70°C.
   23. Let sit at room temperature for 2 minutes.
   24. Centrifuge at maximum speed for 1 minute. Discard the column and keep the microcentrifuge tube with the liquid DNA inside of it.
   25. Store eluted DNA at -20°C.

Data:

After discussing the specifics of the EZNA kit protocol as a class, my group was able to get started with completing our Ludox procedure. However, upon initially centrifuging our samples in both of our 2 mL tubes, we noticed that one of our tubes had clearly pelleted correctly, while the other one did not. We believe that this may have been because of the presence of excess Ludox in our cell layer sample in the 2 mL tube and had to adjust the procedure slightly so that we could get a pellet from that sample, even though it didn’t work properly the first time. With the tube that pelleted correctly, we followed the procedure as stated above, however with the tube that didn’t pellet correctly we had to improvise a bit and revise the procedure so that we could still isolate DNA from this sample. We instead followed this procedure with the tube that didn’t pellet correctly after the first time it was centrifuged:

1. We removed enough of the supernatant solution from the 2 mL tube to fill two 1.5 mL centrifuge tubes to about the halway mark each. Therefore, we had about .75 mL of supernatant solution in each of the 1.5 mL tubes and left the rest in the 2 mL tube.
2. We then added 1000 μL of PBS and filled each 1.5 mL centrifuge tube. We added enough PBS to our 2 mL tube so that it had about the same volume of our tube that was following the normal procedure (at step 3 of the Ludox procedure listed above.)
3. Centrifuge all four tubes at 3000 g for five minutes.
4. At this point, we were able to clearly see that our cells had successfully pelleted in each of the tubes. Remove supernatant from each tube. Be careful not to disturb the pellet.
5. Instead of adding 100 μL of PBS to resuspend the cells in our tubes, we added 50 μL PBS to each tube, so that way we would still be able to combine all of the cell material solution together to get a total of 200 μL of combined solution.
6. From this point on we were able to follow the normal procedure and created the iodine cell solution to observe and count the cells present in our solution.

Because our group had to go through some extra steps and had to follow an alternate procedure, we worked slower and fell behind in completing the entire procedure for both protocols. While we were able to create the iodine-cell solution in our Ludox procedure and were able to observe some of the drops from our sample, we were not able to count the number of cells we found in each drop. While the rest of the class observed five 2 μL drops from the samples, our group was advised to view five 1 μL drops, since the rest of the groups were finding a large number of cells in their drop, making it difficult to count. This is important to note because our calculations may look different than the rest of the class because of this. Even after reducing the size of the drop to 1 μL, we discovered that there were still too many cells to count in our sample drops and we were advised by Dr. Adair to perform a further dilution to accurately count the number of cells in the drop. Therefore, I will come back to lab during open lab tomorrow to perform the dilution and count the number of cells in each drop. While an exact number of cells was not obtained for any of the drops, I was able to observe that there was notable diversity in the morphology of the cells in our drops. The cells were hard to distinguish among the debris, however they appeared to be small, circular brown/black dots.

After we observed the cells in our sample, we began our procedure for the EZNA DNA extraction kit, however we did not get very far into the procedure because there was very little time left of class. We completed up to step 6 (listed above) in the EZNA procedure. Tomorrow I will come back during open lab to complete the rest of this procedure so that we will be caught up and ready for our next steps in the next lab. I labeled our 2 mL tube sample for the EZNA protocol as “Group 7, section 21” and indicated on the side of the tube that we had completed up to step 6. The tube was stored in a yellow rack along with our iodine-cell sample from the Ludox procedure to be observed and counted tomorrow during open lab. The small centrifuge tube of the iodine solution is labeled “Group 7” and is right beside our 2 mL tube in the yellow rack.

Conclusion:

During this lab, my group faced minor setbacks in the procedure that slowed us down and forced us to think of ways that we could correct it in order to achieve results that would be similar and comparable to the rest of the class. This allowed us to experience circumstances that many scientists face when developing a new protocol and by encountering these hurdles in our procedure, we were able to put into practice the creativity needed when encountering a problem that you don’t have the answer to. We needed to push ourselves and develop a way around the problem, and in the end we were able to come to a conclusion that allowed our experiment to get back on track with the rest of the normal procedure. In addition, since we had to go about the procedure a little differently than the rest of the class and we worked at out own pace, we fell behind many of the groups when working on the EZNA protocol. Therefore, future steps would be to complete the rest of this DNA extraction procedure during open lab tomorrow so that our DNA will be extracted and stored for the next lab. Future steps for next week’s lab would be to begin the amplification process with PCR and our specific eukaryotic primers that will target the V4 and v9 region of the 18s rRNA. Also, prior to this we could go upstairs to the lab and shine a UV light on our DNA samples at a certain frequency to prove that there is DNA present in the sample. In conclusion, I am looking forward to the next class and how we will be moving on to examine the DNA we have extracted in future labs.

Purpose:

The purpose of this lab was to be very specific in making our final revisions to our Ludox protocol so that it would be reliable and replicable to other scientists looking for a procedure for DNA isolation and extraction for metabarcoding. Our goal was to make it as specific as possible and then proceed to conduct our experiment and isolate DNA from the same soil sample based on our revised protocol. In addition, the purpose was also to practice our pipetting skills and introduce new lab equipment, such as the serological pipette. We tested the accuracy of the pipettes, as well as our own personal accuracy to ensure that we become confident in these general lab techniques that we will be using often.

Procedure:

1. Listen to lecture on how the different uses of a serological pipette. Prepare to practice taking your own measurements with this kind of pipette by obtaining all the needed materials.
2. Practice pipetting with the serological and micro-pipettes by measuring out different volumes of water and weighing them using a scale and weighing dish. Determine the accuracy of the pipette and your skills by comparing the observed weight of the water you pipette and the expected weight of the water, determined by calculations based on the knowledge that 1 mL of water should weigh 1 gram. Use both the manual and button apparatus for the serological pipette for practice. Complete the following tasks to analyze your pipetting skills:
   • Pipette 1 mL (1000µl) of water using the P1000 micro-pipette.
   • Pipette .1 mL (100µl) of water using the P1000 micropipette.
   • Pipette .1 mL (100 µl total) of water by using the P10 and pipetting 10 µl of water into the weighing dish ten times.
   • Pipette 1 mL of water using the serological pipette. Practice with both the button and manual apparatus.
   • Pipette .1 mL of water using the serological pipette.
3. Record the masses of all of the measurements of water, each time you pipette a new amount of water. Record your observations on the QTM.
4. Complete the new specified procedure for the Ludox protocol that is decided upon in class, based on our answers and provisions from our previous pre-lab submissions. A short summary of the protocol and its changes are as follows:
   1. Collect 5 grams of fresh soil from the class’s soil sample. Screen the soil of large debris before placing it into the weighing dish and into your groups small plastic container.
   2. Add 10 mL of water to the soil. Mix thoroughly for 5-10 minutes until the solution is as homogeneous as possible.
   3. Allow the solution to settle for 1-2 minutes. Transfer 3.68 mL of the mixture to a small glass tube. Add 368 µl of 25% glutaraldehyde to “fix” the sample. Vortex the solution and mix briefly for about 30 seconds to 1 minute.
   4. Add 16mL of Ludox to a clear plastic conical tube. Inject a total of 4 mL of the fixed soil sample into the Ludox using the P1000 micro-pipette at about 2 mL down below the surface of the Ludox.
   5. Layer 2 mL of colored water onto the top of the Ludox. (Create the colored water by adding one drop of dye to a small jar of water.)
   6. Centrifuge the tube at 4300 x g for 15 minutes in a swinging bucket rotor.
   7. Extract a total of 4 mL of organic cell layer from the tube by pipetting 2 mL of the cell layer into two 2 mL tubes each.
   8. Centrifuge the tube 2 mL tubes at 3000 x g for five minutes. Turn and mark the tube so that you know which side the pellet will be angled towards when you are done centrifuging.
   9. Carefully remove the excess supernatant by pipetting it out with a P1000. Be careful not to disturb the pellet.
   10. Add 100 µl of PBS buffer to each 2 mL tube to resuspend each pellet. Flick the tube and pipette up and down to mix and effectively resuspend the pellet. Combine the pellets into the same tube for a total of 200 µl.
   11. Place five 2 µl drops onto a concavity slide and count the cells you see using the 40x lens. Record the number of cells you observe and determine your total of cells/µl. Obtain a class average of these numbers.
   12. Dispose of waste and clean materials properly for next class. Place the ludox tubes in a rack by the sink so that they can be placed in a freezer to freeze for later use.
5. Record any observations and results and complete the QTM.

Data:

When I was practicing my pipetting skills with my lab group at the beginning of the lab, I was able to take the measurements of the mass of each sample of water. While the expected mass of 1 mL of water should be 1 gram and the mass of .1 mL should be .1 grams, this was not necessarily what we were able to measure because our scales couldn’t measure the mass past the tenth place and the pipetting or the scale might not have been accurate or reliable. The measurements that I got for micropipetting 1 mL and .1 mL using the P1000 and .1 mL by using the P10 ten times were:

1 mL- 1.0 grams

.1 mL- 0.0 grams

.1 mL(ten 10 µl) – 0.0 grams

The measurements I recorded for pipetting 1 mL and .1 mL with the serological pipette were:

1 mL- 1.0 grams

.1 mL- 0.0 grams

I was not able to get the accuracy of measuring 0.1 mL of water to result in a mass of 0.1 grams for any of the measurements or different types of pipettes. Instead, the scale would just read that I had 0.0 grams of water.

My group was able to successfully work through and closely follow the steps of the procedure, however due to time constraints, we were only able to complete the protocol up to step 7. We used exactly 5.0 grams of soil to create our soil sample mixture. When adding the glutaraldehyde to fix our sample, we injected 368 µl. However, later in the lab it was discovered that the math in our procedure had been wrong and this was actually too much to put in our sample. The effects of accidentally using more glutaraldehyde than intended could be that the sample becomes too fixed and has toxic negative effects on the cells in our sample, preventing them from emerging from their cysts or making it harder to lyse the cells when we go to extract the DNA. After completing the new Ludox protocol steps 1-5, our sample in the conical tube weighed 40.5 grams before being centrifuged in step 6. When we were able to successfully extract a total of 4 mL of organic cell layer from the conical tube after centrifugation, each 2 mL tube weighed 4.o grams, combined for a total of 8 grams when weighed together. This is as far into the procedure as we got and we had to stop and store them to be centrifuged at a later time. They were stored in a yellow rack next to the group’s tubes across from us, along with all of the other tubes from our class. Each of our tubes used today were labeled “Group 7: SS, LR, KI.”

Conclusion:

In conclusion, I was able to practice my lab techniques with pipetting and I am much more confident in my abilities to accurately and effectively measure out different volumes of a sample with both a micro-pipette and serological pipette. This will be very useful in the future, since both of these pieces of equipment are commonly used in the lab. In addition, I was able to realize just how precise and in depth a procedure has to be in order for it to be standard and replicated by others. You have to take a lot of information and other factors into consideration when you are writing a procedure. It is crucial to be as clear and precise as possible, while still having all of the necessary details and information to make your procedure successful. Since we have performed this Ludox procedure multiple times, we are getting the hang of it and are able to work on smoothing out the kinks that may be causing problems in our procedure and we are one step closer to finalizing our protocol. The next steps we hope to complete in our lab would be to completely finish the procedure and record our observations and results, as well as create a control sample to be used to analyze its results and compare with our own experimental soil samples.