Notebook
From July 31st to August 6th
Experiment Title:
Volatilization curve of formaldehyde (control group experiment)
Introduction:
The goal of this experiment is to test how formaldehyde can volatilize through time as well as observing certain patterns of the curve. This experiment is a precursor of the absorption curve of E. coli in the following experiments.
Experimentation Instruments:
1. blue cap bottle
2. cap with the same size as the probe of the device
3. formaldehyde concentration testing device
4. bottle of saturate 35% concentrated formalin
5. 20-200µL micro pipette
Procedures:
1. Clean the biosafety cabinet; use ventilation, and ultraviolet light techniques for detoxification while setting up the experiment.
2. Turn on the formaldehyde concentration testing device and wait 15min for standardization.
3. Use sealing tape to seal the cap with the probe of the device sticking into the cap.
4. Pipette 20µL of formaldehyde into the bottle and record concentration of formaldehyde in the bottle on the screen every five minutes until the concentration remains stable.
Raw data:
Graph:
Analysis:
According to the data, the stabilized concentration is approximately 6.20mg/m³. Formalin volatized extremely fast and had its steepest curve in the first 5 minutes. Since the final formaldehyde concentration is very concentrated, we may need to dilute formalin and reconduct this experiment with more diluted formalin.
Instructors:
Wenchen Song, Li Cheng
Experiment Conductors:
Zhenhao Yan, Qinming Zhang, Yuelin Lao, Xiaolong Cheng
Experiment Title:
Volatilization curve of formaldehyde (control group experiment)
Introduction:
The goal of this experiment is to test how formaldehyde can volatilize through time as well as observing certain patterns of the curve. This experiment is a precursor of the absorption curve of E. coli in the following experiments.
Experimentation Instruments:
1. 2 blue cap bottles
2. 1 cap with the same size as the probe of the device
3. 1 formaldehyde concentration testing device
4. 1 bottle of saturate 35% concentrated formalin
5. 1 20-200µL micro pipette
6. 20mL of water
7. 4 centrifuge tubes
8. 2 small centrifuge tubes
Procedures:
1. Clean the biosafety cabinet; use ventilation, and ultraviolet light techniques for detoxification while setting up the experiment.
2. Use the micropipette and pipette 100µL of formalin into one of the centrifuge tubes. Pipette another 900µL of water into the same centrifuge tube. Label the centrifuge tube as 10% concentration.
3. Use the micropipette and pipette 10µL of formalin into one of the centrifuge tubes. Pipette another 990µL of water into the same centrifuge tube. Label the centrifuge tube as 1% concentration.
4. Pipette 20µL of the 10% concentration (3.5% formaldehyde) into the 1 mini centrifuge tube. At the same time, place it into the one of the blue cap bottles as fast as possible without spilling. Quickly close the bottle with the cap that is connected to the detection device. Make sure the mini centrifuge tube remains stable. Record all the data until the volatilization is stabilized.
5. Repeat procedure 4 with the 1% concentration (0.35% formaldehyde). Record all data for further analyzation for the control group.
Raw data:
Graph:
Graph:
Analysis:
There are two data that must be analyzed. While both volatilization curve shows a steep rise in the 5 minutes and stabilize in around 20-30 minutes, the less concentrated formalin solution shows a much smaller concentration in the same environment. We did, however, encounter an error since the 10% concentration that we tested today showed a higher result than the 100% concentration yesterday. We hypothesize that this error comes from the improper sealing yester causing the tested concentration higher than the actual concentration. We will test our results again in the future if further time is available. In conclusion, we still have not completed our data since our goal was to maintain the formaldehyde concentration after volatilization below 1 mg/m3. We will be testing lower concentration before out actual experiment to finish the control group experiments.
Instructors:
Wenchen Song, Li Cheng
Experiment Conductors:
Zhenhao Yan, Qinming Zhang, Yuelin Lao, Xiaolong Cheng, Yiqi Liu
Experiment Title:
Volatilization curve of formaldehyde (control group experiment)
Introduction:
The goal of this experiment is to test how formaldehyde can volatilize through time as well as observing certain patterns of the curve. This experiment is a precursor of the absorption curve of E. coli in the following experiments.
Experimentation Instruments:
1. 1 blue cap bottle
2. 1 cap with the same size as the probe of the device
3. 1 formaldehyde concentration testing device
4. 1% concentrated formalin (0.35% formaldehyde) from 08.01
5. 1 20-200µL micro pipette
6. 1 10-1000µL micro pipette
7. 20mL of water
8. 1 empty centrifuge tube
9. 1 small centrifuge tube
Procedures:
1. Clean the biosafety cabinet; use ventilation, and ultraviolet light techniques for detoxification while setting up the experiment.
2. Use the micropipette and pipette 100µL of 1% concentration (0.35% formaldehyde) into one of the centrifuge tubes. Pipette another 900µL of water into the same centrifuge tube. Label the centrifuge tube as 0.1% concentration.
3. Pipette 20µL of the 0.1% concentration (0.035% formaldehyde) into the 1 mini centrifuge tube. At the same time, place it into the one of the blue cap bottles as fast as possible without spilling. Quickly close the bottle with the cap that is connected to the detection device. Make sure the mini centrifuge tube remains stable. Record all the data until the volatilization is stabilized.
Raw data:
Graph:
Analysis:
According to the data, the stabilized concentration is approximately 0.645 mg/m³. Formalin volatized extremely fast and had its steepest curve in the first 10 minutes. On our fourth trial with the 0.1% concentrated formalin, we reached a stabilized environment with only 0.645 mg/m3 of formaldehyde concentration. Without further instructions, this will be the control group experiment that will be based on.
Instructors:
Wenchen Song, Li Cheng
Experiment Conductors:
Zhenhao Yan, Qinming Zhang, Yuelin Lao, Xiaolong Cheng
From August 7th to August 13th
Title of the experiment:
LB Culture creation
Goal of Experiment:
Create LB culture for reservation of future experiments
Method of experimentation:
Mix and Extraction
Materials needed:
1. 1 Glass rod
2. 1 5L beaker
3. 6 Spatulas
4. 2 1L flasks
5. 2 3L graduated cylinders
6. 1 2L beaker
7. 1 100µL-1mL micropipette
8. 1 pH meter • Peptone
9. D-lactose
10. Sodium Chloride
11. Yeast Extract
12. Deionized water
13. 3 Blue capped bottles
14. 1 weight scale
Instructions:
1. Thoroughly clean and sterilize all testing materials before the creation of LB culture. Clean accordingly to steps 1. water, 2. Dishwashing liquid and 3. Deionized water. Make sure that all the clean instruments are labeled.
2. Measure 2.5L of deionized water using the 3 L graduated cylinder. Pour the water into the 5L beaker.
3. Measure 25g of peptone using the weighting scale. While adding it into the beaker and stirring it with deionized water, measure 12.5g of yeast extract and add into the beaker as well. Repeat this process with 25g of Sodium Chloride (NaCl).
4. Stir thoroughly using the glass rod until all solids are dissolved. Use the pH meter to test the pH of the liquid. If the pH is not at 7.0±0.1, adjust accordingly using 10M Sodium Hydroxide (NaOH) until the pH of solution is at 7.0±0.1.
5. Diverse the 2.5L solution into two 1L flasks and one blue capped bottle. After dilution, make sure the flasks and bottles are sealed nicely.
6. Measure 1L of deionized water and pour the water into the 2L beaker.
7. Measure 20g of peptone using the weighting scale; while adding it into the beaker and stirring, measure 10g of D-Lactose and 5g of NaCl and add into the beaker as well. Stir thoroughly until the solids are dissolved.
8. Use the pH meter to test the pH, and if it is not 7.1±0.1, adjust accordingly using 10M NaOH until the pH is steadily at 7.1±0.1
9. Diverse the 1L solution into 2 blue capped bottles. Make sure the caps are sealed nicely to avoid contamination.
10. Clean all flasks, cylinders and beakers using water, soap and deionized water. Repeat this process until all solids have been cleaned off the materials. Gently put them back in designated areas.
11. Make sure the lab bench is cleaned up thoroughly. Leave the lab bench as it was before the experiment.
Issues encountered and How to improve:
• When measuring the material needed for the creation of LB culture, Leo Liu weighed more than he needed.
• Without asking teacher, he dumped the extra material into the waste bin.
• This is not only wasting material, but also may possibly contaminate other materials.
• This is a very serious issue. Next time, we must ask the teacher for instructions and advice whenever we encounter any problems or questions or issues. Like what Mr. Song taught before the first experiment session, don’t do things without getting permission by the teachers.
Experiment Title:
Interlab Calibration 3: Florescence standard curve
Introduction:
The goal of this experiment is to create a standard fluorescence curve for fluorescein concentration. Prepare a dilution series of fluorescein in four replicates and measure the fluorescence in a 96 well plate in the plate reader.
Materials and Experiment Instruments:
1. Fluorescein (provided in kit)
2. 10ml 1X PBS pH 7.4-7.6
3. 96 well plate black with clear flat bottom
4. 20-200μL micropipette
5. 3 small tubes
Procedures:
1. Prepare 10x fluorescein stock solution (100μM) by resuspending fluorescein in 1mL of 1xPBS
2. Dilute the 10x fluorescein stock solution with 1xPBS to make a 1x fluorescein solution with concentration 10μM: 100μL of 10x fluorescein stock into 900μl 1x PBS
3. Add 100μL PBS into wells A2, B2, C2, D2…A12, B12, C12, D12
4. Add 200 μl of fluorescein 1x stock solution into A1, B1, C1, D1
5. Transfer 100 μl of fluorescein stock solution from A1 into A2
6. Mix A2 by pipetting up and down 3x and transfer 100 μl into A3...
7. Mix A3 by pipetting up and down 3x and transfer 100 μl into A4...
8. Mix A4 by pipetting up and down 3x and transfer 100 μl into A5...
9. Mix A5 by pipetting up and down 3x and transfer 100 μl into A6...
10. Mix A6 by pipetting up and down 3x and transfer 100 μl into A7...
11. Mix A7 by pipetting up and down 3x and transfer 100 μl into A8...
12. Mix A8 by pipetting up and down 3x and transfer 100 μl into A9...
13. Mix A9 by pipetting up and down 3x and transfer 100 μl into A10...
14. Mix A10 by pipetting up and down 3x and transfer 100 μl into A11...
15. Mix A11 by pipetting up and down 3x and transfer 100 μl into liquid waste
16. Repeat dilution series for rows B, C, D
17. Measure fluorescence of all samples in instrument
Raw Data:
Fluorescein standard curve
Analysis:
According to the data, as the solution gets more dilute, the value of fluorescence becomes smaller. The highest data is obviously the column one, which was the only fluorescent 1x stock solution and hadn’t been diluted. However, there is one outlier at B3 and B4, but the reason was still unclear. The control group column 12 contains PBS buffer only, so the fluorescence values are very small.
Instructors:
Wenchen Song, Li Cheng
Experiment Conductors:
Yiqi Liu, Qinming Zhang
Experiment Title:
Cell Growth, sampling, and assay
Introduction:
The goal of this experiment is to continue the experiment conducted in Day 2, which was to grow cells (2 colonies from each transformation plate) and to test OD and fluorescence of those overnight cultures. The growth of the cells will be further diluted and analyzed.
Experimentation Instruments:
1. LB+Chloramphenicol solution
2. The 16 samples of cultures that were previously grown overnight
3. mass spectrometer
4. 1 eppendorf tube
5. 2 20-200µL micro pipette+caps
6. 1 Electric Pipette Aid
7. 96 well plate, black with clear flat bottom preferred (provided by team)
Procedures:
1. Make a 1:10 dilution of each overnight culture in LB +Chloramphenicol (10 microliter of culture into 90 microliters of LB +Chloramphenicol)
2. Using the 20-200µL micro pipette, extract 100 μl of each sample and release into each well of the plate according to the diagram below:
3. Measure the Abs600 of these diluted cultures, use a mass spectrometer.
4. Using perhaps an Excel sheet, calculate the amount of LB +Chlor needed to dilute these cultures further into the concentration of 0.02.
5. Take 500 μL samples of the cultures at 6 hours of incubation into 1.5 ml Eppendorf tubes. Place samples on ice.
6. Incubate the remainder of the cultures at 37°C and 220 rpm for 6 hours.
7. Measure the samples again according to the diagram shown under step 2. Record the new data.
8. Record data shown on the Excel sheet.
Raw results:
D at t=0h
A to H in columns 1 to 9 and C to H of column 10 are blanket wells. 10a and 10b are control. Column 11 are number 1 colonies, and column 12 are number 2 colonies.
Fluorescence at t=0
OD at t=6h:
For numbers 1-9:
Negative, Positive, 1, 2, 3, 4, 5, 6, control group (LB +Chlor)
Fluorescence Reading at t=6h
Analysis:
At t=0h:
According to the data, the positive control group’s fluorescence values in this data are larger than those in negative control group, which means that the data can be accepted. The data from colony two of device 5 is much smaller than the colony one of device 5. The highest data is colony 4, which is much larger than the rest of the data. The reason behind this error was unclear, but overall, the whole data has improved.
At t=6h:
The results obtained this time were much more reliable and accurate than the ones previously achieved. There is a big difference between the fluorescence reading of negative and positive control, and the difference of data within each set of variable are very small. The highest could be seen in the fluorescence reading of group 6 (330,337,351,324 for colony 1, and 60,61,65,66 for colony 2), the reasons that attribute to this difference remain partially unclear, but perhaps due to failure in growth of cells of the colony. Overall, the results are a good reflection of the experiment.
Issues encountered today:
One of our overnight culture (1-2 or 1-1) had no proliferation of the colony, discovered this morning at the beginning of dilution. We suppose the reason to be failure of colony selection in yesterday, no colony was picked up and, the tip was empty. We used both cultures from another culture in group one instead of the empty one. The suggested solution can be to touch colonies harder when picking them and slow down the steps to make sure every tip has colony on it.
When adding diluted cultures into 96-well plate, Jerry mistakenly added 2-2 into 3-2’s position. In order to fix his misplacement, we changed positions of 2-2 and 3-2 on the plate, adding 3-2 into initially 2-2’s position. Therefore, when extracting 500 μL of cultures on to the plate, make sure that the order is correct. A preferred method, if the number of people allows, is to have another person to check if the order is correct.
Instructors:
Wenchen Song, Li Cheng
Experiment Conductors:
Zhenhao Yan, Qinming Zhang, Yiqi Liu, Xiaolong Cheng
From September 18th to Septemper 25th
Experiment title:
HindIII EGFP plasmid restriction digestion
Materials:
DNA: 2ul(~0.5ug)
Buffer: 2.5ul
HindIII: 0.5ul
DdH2O: 20ul
Results:
This time the purified concentration of the plasmid wasn’t high enough, only about 16ng/ul.
We would repeat this experiment.
Conductor： YZH
From September 26thth to October 1st
Experiment title:
HindIII EGFP plasmid restriction digestion
Materials:
DNA: 5ul (~2ug)
Buffer: 10ul
HindIII: 2ul
DdH2O: 83ul
Results:
This time the purified concentration was much higher than previous, about 39.5ng/ul.
The recollected parts could be used for following expeirments.
Conductor: WXY
Experiment title:
Gibson Assembly of plasmid pUC57-EGFP and gene FrmR
Description
We used Giblin Assembly method to transfer Nluc into plasmid puc57-EGFP. Length of plasmid puc57-EGFP was 3744 pb while that of FrmR was 876. First we PRCed the two parts. Purified concentration of pUC57-EGFP was 39.5 ng/ul, 0.016 pmol/ul, while that of FrmR was 60.6 ng/ul, 0.105 pmol/ul. And reaction volume for Puc57-EGFP was 2ul, while that of FrmR was 0.915ul.
They were transformed into 50ul of DH5a and cultured overnight under 37 degree.
Conductor: YZH
Experiment title:
Verification of pUC57-EGFP-FrmR assembly by colony PCR
Results:
Primer “a” was the primer of EGFP, which had expected size of 720bp.
Primer “b” was the primer of FrmR, which had expected size of 376bp.
Analysis:
In order to make sure the correctness of our plasmid, we did verification tests. We PCRed primers from plasmid to check. The following is the protocol of this verification test:
Template DNA preparation:
1. pick 5 colonies from transformation plate
2. dissove colony in 30ul of H2O
3. boil it at 98℃ for 10 min ,then put it at room temp for a while
4. pipette 2ul of the supernatant as template for colony PCR
As the gel graph shows, the length of primer a, primer of EGFP, was around 750 bp, and that of primer b, primer of FrmR, was around 400bp. Both of them match with our expectations of primer length. We succeed to construct pUC57-EGFP with gene FrmR!
Conductor： WXY
Experiment title:
Transform plasmid gfa into strain BL21
Because expression of GFA in DH5a wasn’t as well as we expected, we decided to change DH5a into BL21, a kind of frequently-used strain in which GFA may be expressed better and more stably.
First of all, we transformed 3ul of plasmid GFA into 50ul of DH5a and cultured it overnight under 37 celsius
Next: template DNA preparation:
1. pick 3 colonies from transformation plate
2. dissolve colony in 30ul of H2O
3. boil it at 98℃ for 10 min ,then put it at room temp for a while
4. pipette 2ul of the supernatant as template for colony PCR
Results:
Conductor: YZH
Experiment title:
Nluc gel extraction
Results:
Lengths of Nluc matched with the expected size of 516bp.
The final concentration of recollection was 33ng/ul, 39ng/ul
Conductor: YZH
Experiment title:
Submission part PCR
Description
We PCRed all of our parts after extracting and collecting them.
Materials:
1. Template DNA: 1ul (-20ng)
2. 5x Buffer: 10ul
3. dNTP: 1ul
4. Q5 enzyme: 0.5ul
5. DdH2O: 32.5ul
Notice; The volume of FrmR WAS 0.5 ul (~30 ng)
The extension time was 45s, 35cycles.
The results:
Expected size for Nluc, pfrmr, Frmr, EGFP, and GFA were 516, 200, 376, 720, and 585bp. For each parts we repeated PCR twice. Purified concentrations for Nluc were 77.7 and 75.8 ng/ul; that for pFrmR was 15.5; that for EGFP were 37.8 and 42.1 ng/ul; and that for GFA were 68 and 68.8 ng/ul. As the graph showed, digestion recovery of FrmR wasn’t shown clearly.
Conductor: CXL
Experiment title:
EcoR-HF&PstI submission part restriction digestion
Materials:
(1)Digestion of Nluc:
1. Buffer: 4ul
2. EcoRI-HF: 2ul
3. Pstl: 2ul
4. DNA: 26ul(-2ug)
5. DdH20: 38ul
(2)Digestion of EGFP:
1. Buffer: 4ul
2. EcoRI-HF: 2ul
3. Pstl: 2ul
4. DNA:48ul(-2ug)
5. DdH20: 16ul
(3)Digestion of GFA:
1. Buffer: 4ul
2. EcoRI-HF: 2ul
3. Pstl: 2ul
4. DNA: 30ul(-2ug)
5. DdH20: 34ul
(4)Digestion of pFrmR:
1. Buffer: 4ul
2. EcoRI-HF: 2ul
3. Pstl: 2ul
4. DNA: 28ul(-0.4ug)
5. DdH20: 4ul
Results:
In this step the enzyme digestion recovery concentration for Nluc, EGFP, GFA, and pFrmR are 38.8, 49.4, 55.4, and 4.4 (ng/ul)
Conductor: YZH
Experiment title:
Submission part ligatio
Materials:
(1)Ligation of Nluc:
1. 10x buffer: 2ul
2. T4 DNA ligase: 1ul
3. Vector DNA: 5ul(~25ng, 0.020pmol)
4. DNA: 0.5ul(~0.060pmol)
DdH2O:11.5ul
(2)Digestion of EGFP:
1. 10x buffer: 2ul
2. T4 DNA ligase: 1ul
3. Vector DNA: 5ul(~25ng, 0.020pmol)
4. DNA: 0.5ul(~0.060pmol)
5. DdH2O:11.5ul
(3)Ligation of GFA:
1. 10x buffer: 2ul
2. T4 DNA ligase: 1ul
3. Vector DNA: 5ul(~25ng, 0.020pmol)
4. DNA: 0.5ul(~0.060pmol)
DdH2O:11.5ul
(4)Ligation of pFrmR:
1. 10x buffer: 2ul
2. T4 DNA ligase: 1ul
3. Vector DNA: 5ul(~25ng, 0.020pmol)
4. DNA: 3ul(~0.060pmol)
5. DdH2O:9ul
The process lasted for 1h or 2h. All the genes are transferred into E. coli DH5a, and bacteria were cultured under 39 degree for 2 days.
Results:
At least on gel plate every parts had successfully ligated parts. Next step will be the verification test to make sure we transformed parts into DH5a correctly.
Conductor: WXY
Experiment title:
Verification of submission part ligation by colony PCR
Description:
After ligating our parts, we did verification tests to make sure that our parts are successfully transformed into DH5a.
Materials:
1. Tag mix: 6ul
2. DNA:2ul
3. Primer F/R: 1ul
4. H2O: 2ul
Procedure:
Template DNA preparation:
1. pick colonies from each plate
2. dissove colony in 30ul of H2O
3. boil it at 98℃ for 10 min ,then put it at room temp for a while
4. pipette 2ul of the supernatant as template for colony PCR
Results:
As the gel graph showed, all of our parts were in length of their expected size, and there wasn’t much fluctuations.
Conductor: CXL
