Pipetting Technique and Accuracy Evaluation

Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:40 UTC 05 Jan 2025 20:24:54 UTC Pipetting Technique and Accuracy Evaluation Project: Molecular biology lab book Author: Entry Created On: 15 Sep 2024 02:20:39 UTC Entry Last Modified: 26 Sep 2024 01:25:11 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC SATURDAY, 2024-09-14 Purpose By using micro-pipettor and serological pipettes, understand and practice pipetting skill. The accuracy when using pipette is critical aspect for carrying out successful labatory experiment. From this protocol, it is able to carryout proper pipetting technique as well as calculating percent error of pipetting accuracy. Biology 2290 Pipetting In this lab you will get to practise using both a micro-pipettor and a serological pipette. Accurate pipetting skills are obtained through practice and using proper technique. This is a fundamental skill for a variety of science disciplines. Serological Pipetting Exercise: 1. Weigh the empty 100 ml beaker and record this in your benchling. 2. Fill second beaker with water from white tap 3. Using a 5 mL serological pipette, individually pipette the following volumes into the beaker: a. 0.5 mL (500ul) b. 1.0 mL c. 2 mL d. 3 mL e. 3.5 mL 4. The combined volume should be 10 mL. The density of water at room temperature is 0.9976 g/mL. Therefore, your 10 mL should weigh 9.98g. Weight of empty beaker =53.88g Weight of beaker + water =63.538 Weight of water only =9.65 1. Now we will calculate the % error in your pipetting using the following equation: Measured weight of water – Calculated weight of water X 100 =(10-9.65)=3.61 Calculated weight of water An % error of 1-2% is not unusual. If your % error is greater than 5% you should review your pipetting technique. Micro-Pipetting Exercise Be careful to use the correct pipette. Use water from white tap. You don’t have to change tips for these exercises. As you do these exercises, look at the pipette tip when it is holding a volume so you can learn to check your pipetting visually. P1000 (100/200-1000 µl) – 7 eppendorf (1.5mL microcentrifuge tubes, they have a lot of names) tubes: 1. Add 1000 µl to three eppendorf tubes. 2. Visually check that each tube has the same volume. 3. Remove 750 µl from each tube and pipette into three new tubes. 4. Visually check that each tube has the same volume. On this page: Entry Page Page1 1ofof34 3 Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:40 UTC 05 Jan 2025 20:24:54 UTC 5. Remove 250 µl from the original three and pipette into one new tube. ● Remove 750 µl from this tube – read steps 7-9 before discarding liquid and tip ● How accurate were you? Is some liquid left in the tube? (no liqud are left in the original tube) ● Measure it by drawing it up, and adjusting the pipette volume until all air below the liquid is expelled from the tip. Remember to hold the pipette vertically while doing so. ● Did you draw air up with the last removal of liquid? Measure this volume by adjusting the volume downward until the air is gone and liquid is at the bottom of the tip. Remember to hold the pipette vertically while doing so. (There were no air drawn up) ● The pipettes have an accuracy and precision of about 1% - any more than that is human pipetting error! If you got it very wrong, try again. P200 (20-200 µl) – 3 of PCR 8 well strip tubes, and 1 eppendorf tube: 1. Add 100 µl to each well of a set of strip tubes. 2. Visually check that each tube has the same volume. 3. Remove 50 µl from each tube and pipette into a new set of strip tubes. 4. Visually check that each tube has the same volume. 5. Remove 30 µl from the original strip tubes and pipette into a new set of strip tubes. 6. Visually check that each tube has the same volume. 7. Remove 20 µl from each well of the original strip tubes and pipette(pool) into one new effendorf tube. ● Remove 160 µl from this tube - read steps 9-11 before discarding liquid and tip ● How accurate were you? Is some liquid left in the tube? (There were no liquid left in the tube) ● Measure it by drawing it up, and adjusting the pipette volume until all air below the liquid is expelled from the tip. Remember to hold the pipette vertically while doing so. ● Did you draw air up with the last removal of liquid? Measure this volume by adjusting the volume downward until the air is gone and liquid is at the bottom of the tip. Remember to hold the pipette vertically while doing so. none of the air was drawn up ● The pipettes have an accuracy and precision of about 1% - any more than that is human pipetting error! If you got it very wrong, try again. ● Shake water out of the strip tubes and use in next section P20 (2-20 µl) Larger volume– 1 strip tube and 1 eppendorf tube: 1. Add 18 µl to each well of a set of strip tubes. 2. Visually check that each tube has the same volume. 3. Remove 12 µl from each well and pipette into one new Eppendorf tube. ● Remove 96 µl from this tube - read steps 5-7 before discarding liquid and tip ● How accurate were you? Is some liquid left in the tube? None of the liquid was drawn up ● Measure it by drawing it up, and adjusting the pipette volume until all air below the liquid is expelled from the tip. Remember to hold the pipette vertically while doing so. (There were no liquid left in the original strip tube) ● Did you draw air up with the last removal of liquid? Measure this volume by adjusting the volume downward until the air is gone and liquid is at the bottom of the tip. Remember to hold the pipette vertically while doing so.(at final, there were no liquid drawn up) No liquid was drawn up ● The pipettes have an accuracy and precision of about 1% - any more than that is human pipetting error! If you got it very wrong, try again P20 (2-20 µl) Smaller Volume – 1 set of strip tubes: 1. Add 7 µl to three 0.2 mL (strip) tubes. 2. Visually check that each tube has the same volume. On this page: Entry Page Page2 2ofof34 3 Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:40 UTC 05 Jan 2025 20:24:54 UTC 3. Add 2 µl to each tube of the three tubes with 7 µl . 4. Visually check that each tube has the same volume. 5. Remove 9 µl from each tube and pipette into three new 0.2 mL tubes. 6. Visually check that each tube has the same volume. They all had same volume On this page: Entry Page Page3 3ofof34 3 Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:37 UTC 05 Jan 2025 20:24:54 UTC Restriction Digest and Ligation of Plasmid DNA Using EcoRI and T4 DNA Ligase for Cloning Project: Molecular biology lab book Author: Kyuwon Jung Entry Created On: 15 Sep 2024 02:23:08 UTC Entry Last Modified: 23 Oct 2024 20:57:44 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC SATURDAY, 2024-09-14 Purpose To perform restriction digest on plasmid by using EcoRI during the preparation of cloning. To enable the ligation of insert for subsequent transfromation, he plasmid will be digested Plasmid information Plasmid name: pUC19 Concentration: 25 ng/μL Size: 2686 bp Restriction Digest Kit name: FastDigest, From: Thermofisher 1. Work in partners.As a pair you will complete the following reactions 2. Get three 1.5 mL microcentrifuge tubes and label them “Insert”, “H20”, and “TE” and your initials 3. Look at the three tables below and add the appropriate components to each tube: a. You will first have to calculate the volume of plasmid to add to achieve 50 ng. Insert Tube Table1 A B 1 Component: Insert tube Volume 2 Autoclaved dH2O 4µL 3 5x FastDigest Buffer 2 µL 4 50 ng Plasmid DNA. 2µL 5 EcoRI 2 µL 6 TOTAL 10µL H2O Tube On this page: Entry Page Page4 1ofof34 8 Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:37 UTC 05 Jan 2025 20:24:54 UTC Table2 A B 1 Component: H20 tube Volume 2 Autoclaved dH2O 4µL 3 5x FastDigest Buffer 2µL 4 50 ng Plasmid DNA. 2µL 5 EcoRI 2 µL 6 TOTAL 10 µL TE Tube Table3 A B 1 Component: TE tube Volume 2 Autoclaved dH2O 4µL 3 5x FastDigest Buffer 2 µL 4 TE buffer 2µL 5 EcoRI 2 µL 6 TOTAL 10 µL 1. Gently mix the tube by pipetting up and down. If there is liquid on the sides of the tube give it a quick spin in the centrifuge. 2. Place tube in 37 °C water bath for 15 min. 3. Return tubes to ice Ligation Protocol Kit name: T4 DNA ligase, From: Thermofisher 1. You are performing a ligation reaction with an insert:vector of 3:1. You will need to calculate the volume of insert to add to your tube. The ng of vector (plasmid) is kept constant at 50 ng. 2. To your restriction digest tubes, add the following: Ligation with insert On this page: Entry Page Page5 2ofof34 8 Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:37 UTC 05 Jan 2025 20:24:54 UTC Table4 A B 1 Component Volume 2 5X Ligase reaction buffer 4 mL 3 Plasmid DNA (in tube already) 10 μL 4 Insert DNA. 1.12μL 5 Autoclaved, dH2O 2.88μL 6 T4 DNA ligase 2μL 7 TOTAL 20 μL Ligation with H20 Table5 A B 1 Component Volume 2 5X Ligase reaction buffer 4 µL 3 Plasmid DNA (in tube already) 10 µL 5 Autoclaved, dH2O 4µL 6 T4 DNA ligase 2 µL 7 TOTAL 20 µL 4 Ligation with TE On this page: Entry Page Page6 3ofof34 8 Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:37 UTC 05 Jan 2025 20:24:54 UTC Table6 A B 1 Component Volume 2 5X Ligase reaction buffer 4 µL 3 TE buffer (in tube already) 10 µL 5 Autoclaved, dH2O 4µL 6 T4 DNA ligase 2 µL 7 TOTAL 20 µL 4 1. Gently mix by pipetting up and down and quick spin in the centrifuge if necessary. 2. Incubate both tubes at room temperature for 10 min (In the tube rack on your bench). 3. Once done, place in tube rack at front of room. Result Table7 Plates Colony count 1 TE 10^-6 226 white collonies 2 TE 0 Collonies 3 H2O 49 blue collonies 4 Insert 24 white 26 blue collonies 5 Plasmid 256 Blue Collonies On this page: Entry Page Page7 4ofof34 8 Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:37 UTC 05 Jan 2025 20:24:54 UTC Ligation Protocol WITH T4 DNA Ligase (M0202) Introduction Please see the NEB website for supporting information on this protocol. Materials › 10X T4 DNA Ligase Reaction Buffer › T4 DNA Ligase › Vector DNA (4kb) › Insert DNA (1kb) › Nuclease-free water Procedure Set up the T4 DNA Ligase Reaction Note: T4 DNA Ligase should be added last. The table shows a ligation using a molar ratio of 1:3 vector to insert for the indicated DNA sizes. Use NEB calculator to calculate molar ratios. 1. Thaw the T4 DNA Ligase Buffer and resuspend at room temperature. Tip: Alicuote the 10x buffer less concentrated so when thawing, the DTT gets soluble more easily. 2. Set up the following reaction in a microcentrifuge tube on ice: Table1 A B 1 Component Volume (μl) 2 10X T4 DNA Ligase Buffer 3 Vector DNA: 50 ng (0.020 pmol) 4 Insert DNA: 37.5 ng (0.060 pmol) 5 Nuclease-free water 6 T4 DNA Ligase 7 Total 2 17 1 20 3. Gently mix the reaction by pipetting up and down and microfuge briefly. On this page: Entry Page Page8 5ofof34 8 Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:37 UTC 05 Jan 2025 20:24:54 UTC 4. For cohesive (sticky) ends, incubate at 16°C overnight or room temperature for 10 minutes. For blunt ends or single base overhangs, incubate at 16°C overnight or room temperature for 2 hours. 5. Heat inactivate at 65 degrees C for 10 minutes. 00:10:00 6. Chill on ice and transform 1-5 μl of the reaction into 50 μl competent cells. Use 25 uL DH5α cells, and add 2 uL of reaction mixture. On this page: Entry Page Page9 6ofof34 8 Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:37 UTC 05 Jan 2025 20:24:54 UTC Ligation Protocol WITH T4 DNA Ligase (M0202) Introduction Please see the NEB website for supporting information on this protocol. Materials › 10X T4 DNA Ligase Reaction Buffer › T4 DNA Ligase › Vector DNA (4kb) › Insert DNA (1kb) › Nuclease-free water Procedure Set up the T4 DNA Ligase Reaction Note: T4 DNA Ligase should be added last. The table shows a ligation using a molar ratio of 1:3 vector to insert for the indicated DNA sizes. Use NEB calculator to calculate molar ratios. 1. Thaw the T4 DNA Ligase Buffer and resuspend at room temperature. Tip: Alicuote the 10x buffer less concentrated so when thawing, the DTT gets soluble more easily. 2. Set up the following reaction in a microcentrifuge tube on ice: Table1 A B 1 Component Volume (μl) 2 10X T4 DNA Ligase Buffer 3 Vector DNA: 50 ng (0.020 pmol) 4 Insert DNA: 37.5 ng (0.060 pmol) 5 Nuclease-free water 6 T4 DNA Ligase 7 Total 2 17 1 20 3. Gently mix the reaction by pipetting up and down and microfuge briefly. On this page: Entry Page 10 7ofof348 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:37 UTC 05 Jan 2025 20:24:54 UTC 4. For cohesive (sticky) ends, incubate at 16°C overnight or room temperature for 10 minutes. For blunt ends or single base overhangs, incubate at 16°C overnight or room temperature for 2 hours. 5. Heat inactivate at 65 degrees C for 10 minutes. 00:10:00 6. Chill on ice and transform 1-5 μl of the reaction into 50 μl competent cells. Use 25 uL DH5α cells, and add 2 uL of reaction mixture. On this page: Entry Page 11 8ofof348 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:49:01 UTC 05 Jan 2025 20:24:54 UTC Restriction Digest Protocol for Plasmid and Insert DNA Preparation Project: Molecular biology lab book Author: Kyuwon Jung Entry Created On: 18 Sep 2024 00:40:13 UTC Entry Last Modified: 19 Sep 2024 13:57:28 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC TUESDAY, 2024-09-17 Purpose By performing a restriction digestion followed by a ligation reaction. The restriction digest will cut a plasmid using the restriction enzyme EcoRI to provide suitable ends for ligation. In the ligation stage, the digested plasmid is combined with an insert in a 3:1 ratio to form a recombinant DNA construct. Controls containing water and TE buffer are included to ensure that the ligation reaction is specific and efficient. Restriction Digest Kit name: FastDigest, From: Thermofisher 1. Work in partners.As a pair you will complete the following reactions 2. Get three 1.5 mL microcentrifuge tubes and label them “Insert”, “H20”, and “TE” and your initials 3. Look at the three tables below and add the appropriate components to each tube: a. You will first have to calculate the volume of plasmid to add to achieve 50 ng. Table1 A B 1 Component: Insert tube Volume 2 Autoclaved dH2O 4 µL 3 5x FastDigest Buffer 2 µL 4 50 ng Plasmid DNA. 2 µL 5 EcoRI 2 µL 6 TOTAL 10 µL On this page: Entry Page 12 1ofof344 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:49:01 UTC 05 Jan 2025 20:24:54 UTC Table2 A B 1 Component: H20 tube Volume 2 Autoclaved dH2O 4µL 3 5x FastDigest Buffer 2 µL 4 50 ng Plasmid DNA. 2µL 5 EcoRI 2 µL 6 TOTAL 10 µL Table3 A B 1 Component: TE tube Volume 2 Autoclaved dH2O 4µL 3 5x FastDigest Buffer 2 µL 4 TE buffer 2µL 5 EcoRI 2 µL 6 TOTAL 10 µL 1. Gently mix the tube by pipetting up and down. If there is liquid on the sides of the tube give it a quick spin in the centrifuge. 2. Place tube in 37 °C water bath for 15 min. 3. Return tubes to ice Ligation Protocol Kit name: T4 DNA ligase, From: Thermofisher 1. You are performing a ligation reaction with an insert:vector of 3:1. You will need to calculate the volume of insert to add to your tube. The ng of vector (plasmid) is kept constant at 50 ng. 2. To your restriction digest tubes, add the following: Ligation with insert On this page: Entry Page 13 2ofof344 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:49:01 UTC 05 Jan 2025 20:24:54 UTC Table4 A B 1 Component Volume 2 5X Ligase reaction buffer 4 µL 3 Plasmid DNA (in tube already) 10 µL 4 Insert DNA. 2µL 5 Autoclaved, dH2O 2µL 6 T4 DNA ligase 2 µL 7 TOTAL 20 µL Ligation with H20 Table5 A B 1 Component Volume 2 5X Ligase reaction buffer 4 µL 3 Plasmid DNA (in tube already) 10 µL 5 Autoclaved, dH2O 4µL 6 T4 DNA ligase 2 µL 7 TOTAL 20 µL 4 Ligation with TE On this page: Entry Page 14 3ofof344 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:49:01 UTC 05 Jan 2025 20:24:54 UTC Table6 A B 1 Component Volume 2 5X Ligase reaction buffer 4 µL 3 TE buffer (in tube already) 10 µL 5 Autoclaved, dH2O 4µL 6 T4 DNA ligase 2 µL 7 TOTAL 20 µL 4 1. Gently mix by pipetting up and down and quick spin in the centrifuge if necessary. 2. Incubate both tubes at room temperature for 10 min (In the tube rack on your bench). 3. Once done, place in tube rack at front of room. On this page: Entry Page 15 4ofof344 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:49 UTC 05 Jan 2025 20:24:54 UTC Transformation of pAMP into E. coli JM101 Project: Molecular biology lab book Author: Kyuwon Jung Entry Created On: 23 Sep 2024 17:24:07 UTC Entry Last Modified: 23 Sep 2024 18:08:39 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC MONDAY, 2024-09-23 Purpose Transform pAMP into E. coli JM101 cells using a ligation reaction. The goal is to compare transformation efficiency under different conditions: plasmid, H₂O control, and TE buffer. Transformation Protocol (Not a kit) 1. Collect 4 tubes of competent E.coli JM101 (250 uL in each tube) in your ice bucket. ALL steps must be performed on ice. Label one tube” plasmid”, one tube “TE”, one tube “insert”, and one tube” H20” and Put your initials on all tubes. 2. Add 10 uL of ligation with insert to the tube of cells labelled insert 3. Add 10uL of ligation with H20 to the tube of cells labelled H20 4. Add 10 uL of ligation with TE buffer to the tube of cells labelled TE 5. Add 2 uL pAMP to the tube of cells labelled plasmid. 6. Mix gently by flicking the tube. 7. Incubate on ice for 5 min. 8. Incubate in 45 °C water bath for 100 s 9. Incubate on ice for 1 min 10. Add 250 uL LB broth to each tube 11. Incubate in 37 °C water bath for 15 min. 12. After 15 min incubation tubes can be kept at room temperature. 13. Gently mix your tubes by flicking or pipetting up and down to make sure cells have not settled. 14. Using your TE tube, make a 10-6 dilution. Dilute using the 0.5% NaCl “dilution solution”. Once your dilution is complete, keep the original buffer tube and your 10-6 dilution tube. All other tubes can be disposed of (10-2, 10-4). 15. You should now have 5 tubes: Insert, H20, Plasmid, TE and 10-6 dilution of TE. 16. Tidy your work area so nothing flammable is out. Collect four LB+100 ug/mL ampicillin plates (pink) and one LB plate (yellow). Label them around the circumference with your name, section number and what is on the plate (see step 18 for this information). 17. Light your Bunsen burner 18. Using sterile technique, one plate at a time, add 100 uL of below and spread: a. Insert on LB + amp plate b. H20 on LB + amp plates c. Plasmid on LB + amp plate d. TE on LB + amp plate e. 10-6 dilution of TE on LB plate 19. Turn off the Bunsen burner. 20. Gather your 5 plates and put an elastic around them. 21. Place them in the wire rack upside down (label side up). On this page: Entry Page 16 1ofof342 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:49 UTC 05 Jan 2025 20:24:54 UTC 22. They will be incubated overnight at 37 °C. 23. Lysol your bench 24. Wash your hands. On this page: Entry Page 17 2ofof342 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:55 UTC 05 Jan 2025 20:24:54 UTC Preparation and pH Adjustment of Tris Buffer Project: Molecular biology lab book Author: Kyuwon Jung Entry Created On: 25 Sep 2024 18:06:31 UTC Entry Last Modified: 26 Sep 2024 14:01:27 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC WEDNESDAY, 2024-09-25 Purpose The purpose of this experiment is to prepare a Tris buffer solution, essential for maintaining pH stability during biological experiments such as DNA electrophoresis. We will create a buffer with a final concentration of 50 mM Tris and 100 mM NaCl, adjusted to pH 7.5 using HCl. This pH is optimal for many enzymatic reactions and helps prevent degradation of nucleic acids, ensuring reliable results in molecular biology applications. pHing In Biology Protocol You will be working in pairs to make a Tris buffer (part of TBE). The pH meters have already been calibrated and are ready to use. 1. Using a weigh boat and the “Tris” scoopula, weigh out 0.606g of Tris and then dump it in the provided beaker. 2. Using a weigh boat and the “NaCl” scoopula, weigh out 0.584xszg of NaCl and add it to the beaker. 3. Using a graduated cylinder, measure out 60 mL of H2O and add this to the beaker. 4. Place a magnetic stir bar in the beaker and place on the stir plate. Stir mixture until completely dissolved. 5. Remove the pH probe from the neutral storage buffer and rinse with a squeeze bottle of water then place in the Tris solution (while it is still stirring – make sure probe is not being hit by stir bar). 6. Observe the pH reading and when the reading stabilizes note the pH. 7. Using the serological pipette slowly (approximately 2 mL at a time) add HCL to the solution while still stirring. Keep track of roughly how much HCL in total you are adding.18.5ml used 8. Add HCL until pH is 7.5. 9. Pour the buffer back into the graduated cylinder and top up volume to 100 mL. 10. Dispose of your buffer in the sink and rinse your beaker before placing it in the bin for glass washing. 18.5ml of HCl was used to make pH to 7.5 To top off volume of 100ml we have used 21.5ml of distilled water The initial pH was 10.40 2 On this page: Entry Page 18 1ofof341 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:46 UTC 05 Jan 2025 20:24:54 UTC Yeast Cell Counting and Viability Assessment Using a Haemocytometer Project: Molecular biology lab book Author: Kyuwon Jung Entry Created On: 25 Sep 2024 18:10:42 UTC Entry Last Modified: 03 Oct 2024 07:56:26 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC WEDNESDAY, 2024-09-25 Purpose The purpose of this lab is to precisely count the number of yeast cells in a culture and measure cell viability with a hemocytometer and trypan blue staining. This experiment teaches essential cell counting, microscopy, and viability assessment techniques by distinguishing between live (clear) and dead (blue) cells, which are critical for a variety of biological and biotechnological applications such as culture density measurements and cell health assessments. Preparation of Haemocytometer: 1. Give the tube of yeast a gentle flick to resuspend cells (they tend to settle on the bottom of the tube) 2. Pipette 25 uL of yeast into a new 1.5 mL tube 3. Add 25 uL of trypan blue to this tube of yeast and pipette up and down to mix 4. Place cover slip on haemocytometer 5. Set your pipette to 10 uL and draw up the yeast and trypan blue mixture 6. Carefully begin loading one side of the Haemocytometer in the notch. Once first chamber is full, repeat procedure for second chamber. 7. Let sit for 2 min. to allow cells to settle Counting Yeast: 1. Move the objective lenses to the highest position using course adjustment knob 2. Rotate nose piece so 10x objective is in place 3. Place the Haemocytomer on the stage and secure with clips 4. Adjust the stage position so you can see the grids 5. Using the course adjustment knob, slowly lower the objectives towards the Haemocytometer 6. Once in focus, - if you want - carefully rotate nose piece to select for the 40x objective. NOTE: this will be very close to the slide cover so move slowly, use only the fine adjustment knob at this point. Note, you can count under either 10X or 40X objective. 7. Using the stage adjustment knob, centre your first square for counting (one of the four corners or middle) and begin counting yeast cells 8. Record the number of blue cells and the number of clear cells in your lab book 9. Repeat for the four other squares in the chamber and then repeat again for the other chamber.1st chamber (5x5Blue cells: 7 transparent 20) (4x4 16 alive, 4 dead)(19 alive 6 dead)(18 alive 8 dead)(19 alive 8 dead)totalalive= total dead=33 2nd chamber(5x5 17 alive 12dead 4x4 24 alive 12 dead)(15 alive 18 dead)(25 alive 11 dead)(23 alive 10 dead) 10. When done, spray a kimwipe with ethanol and carefully wipe both the cover slip and haemocytometer and return them to the storage boxes 11. Calculate the average number of live cells (clear) in the provided yeast culture and record in your lab book 12. Calculate the cell viability in the yeast culture and record in your lab book. On this page: Entry Page 19 1ofof342 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:46 UTC 05 Jan 2025 20:24:54 UTC Table1 Quadrant Aliv e Dea d 1 5X5 20 7 2 Top right 4X4 16 4 3 Top left 4X4 19 6 4 Bottom right 4X4 18 8 5 Bottom left 4X4 19 8 6 Total 92 33 Table2 Quadrant Aliv e Dea d 1 5X5 17 12 2 Top right 4X4 24 12 3 Top left 4X4 15 18 4 Bottom right 4X4 25 11 5 Bottom left 4X4 23 10 6 Total 104 51 Average number of alive cells in 10 quadrant= 19.6 Cell count= 19.6*250*2=9.8*103 Cell viability= 1-(4.2*103/1.4*104)=0.7 Therefore the cell viability is 70% On this page: Entry Page 20 2ofof342 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:52 UTC 05 Jan 2025 20:24:54 UTC Copy of Yeast Cell Counting and Viability Assessment Using a Haemocytometer Project: Molecular biology lab book Author: Kyuwon Jung Entry Created On: 04 Nov 2024 00:47:37 UTC Entry Last Modified: 04 Nov 2024 00:47:37 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC THURSDAY, 2024-09-26 Purpose The purpose of this experiment is to prepare a Tris buffer solution, essential for maintaining pH stability during biological experiments such as DNA electrophoresis. We will create a buffer with a final concentration of 50 mM Tris and 100 mM NaCl, adjusted to pH 7.5 using HCl. This pH is optimal for many enzymatic reactions and helps prevent degradation of nucleic acids, ensuring reliable results in molecular biology applications. pHing In Biology Protocol You will be working in pairs to make a Tris buffer (part of TBE). The pH meters have already been calibrated and are ready to use. 1. Using a weigh boat and the “Tris” scoopula, weigh out 0.606g of Tris and then dump it in the provided beaker. 2. Using a weigh boat and the “NaCl” scoopula, weigh out 0.584g of NaCl and add it to the beaker. 3. Using a graduated cylinder, measure out 60 mL of H2O and add this to the beaker. 4. Place a magnetic stir bar in the beaker and place on the stir plate. Stir mixture until completely dissolved. 5. Remove the pH probe from the neutral storage buffer and rinse with a squeeze bottle of water then place in the Tris solution (while it is still stirring – make sure probe is not being hit by stir bar). 6. Observe the pH reading and when the reading stabilizes note the pH. 7. Using the serological pipette slowly (approximately 2 mL at a time) add HCL to the solution while still stirring. Keep track of roughly how much HCL in total you are adding.18.5ml used 8. Add HCL until pH is 7.5. 9. Pour the buffer back into the graduated cylinder and top up volume to 100 mL. 10. Dispose of your buffer in the sink and rinse your beaker before placing it in the bin for glass washing. On this page: Entry Page 21 1ofof341 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:44 UTC 05 Jan 2025 20:24:54 UTC E. coli Transformation with Ligation Reactions for Plasmid Insertion Project: Molecular biology lab book Author: Kyuwon Jung Entry Created On: 02 Oct 2024 03:48:19 UTC Entry Last Modified: 02 Oct 2024 05:27:36 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC TUESDAY, 2024-10-01 Purpose The purpose of this experiment is to transform competent E. coli JM101 cells using different ligation reactions to assess the efficiency of plasmid insertion and the viability of cells. By comparing transformation with ligation products containing inserts, TE buffer, H2O, and plasmids, the experiment aims to identify successful incorporation of plasmids through colony growth on LB plates with ampicillin. The experiment also involves making a 10⁻⁶ dilution of the TE buffer sample to quantify transformation efficiency on non-selective media. Procedure 1. Collect 4 tubes of competent E.coli JM101 (250 uL in each tube) in your ice bucket. ALL steps must be performed on ice. Label one tube” plasmid”, one tube “TE”, one tube “insert”, and one tube” H20” and Put your initials on all tubes. 2. Add 10 uL of ligation with insert to the tube of cells labelled insert 3. Add 10uL of ligation with H20 to the tube of cells labelled H20 4. Add 10 uL of ligation with TE buffer to the tube of cells labelled TE 5. Add 2 uL pAMP to the tube of cells labelled plasmid. 6. Mix gently by flicking the tube. 7. Incubate on ice for 5 min. 8. Incubate in 45 °C water bath for 100 s 9. Incubate on ice for 1 min 10. Add 250 uL LB broth to each tube 11. Incubate in 37 °C water bath for 15 min. 12. After 15 min incubation tubes can be kept at room temperature. 13. Gently mix your tubes by flicking or pipetting up and down to make sure cells have not settled. 14. Using your TE tube, make a 10-6 dilution. Dilute using the 0.5% NaCl “dilution solution”. Once your dilution is complete, keep the original buffer tube and your 10-6 dilution tube. All other tubes can be disposed of (10-2, 10-4). 15. You should now have 5 tubes: Insert, H20, Plasmid, TE and 10-6 dilution of TE. 16. Tidy your work area so nothing flammable is out. Collect four LB+100 ug/mL ampicillin plates (pink) and one LB plate (yellow). Label them around the circumference with your name, section number and what is on the plate (see step 18 for this information). 17. Light your Bunsen burner 18. Using sterile technique, one plate at a time, add 100 uL of below and spread: a. Insert on LB + amp plate b. H20 on LB + amp plates c. Plasmid on LB + amp plate d. TE on LB + amp plate On this page: Entry Page 22 1ofof342 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:44 UTC 05 Jan 2025 20:24:54 UTC e. 10-6 dilution of TE on LB plate 19. Turn off the Bunsen burner. 20. Gather your 5 plates and put an elastic around them. 21. Place them in the wire rack upside down (label side up). 22. They will be incubated overnight at 37 °C. 23. Lysol your bench 24. Wash your hands. On this page: Entry Page 23 2ofof342 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:39 UTC 05 Jan 2025 20:24:54 UTC Standard Curve and Insulin Quantification in Spent Media using Colorimetric Assay Project: Molecular biology lab book Author: Kyuwon Jung Entry Created On: 09 Oct 2024 20:20:49 UTC Entry Last Modified: 23 Oct 2024 11:26:35 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC WEDNESDAY, 2024-10-09 Purpose The purpose of this experiment is to create a standard curve for insulin concentration using a serial dilution of an insulin standard and a colorimetric assay. The standard curve will then be used to quantify the insulin concentration in two spent media samples: one experimental sample treated with insulin and one control sample grown with an empty vector. The absorbance of the samples will be measured using both a Spec 20 and a microplate reader to ensure accurate concentration determination. In this lab you will be performing a serial dilution in partners to create a standard curve and then use that curve to determine the concentrations of our treated spent media (experimental sample) and spent media grown with an empty vector (no insert) - this is a control. This standard curve will use a colorimetric assay to determine the concentration. 1. Specs will be turned on already so that they are warmed up. They will be set to 425 nm which is the lambda max for the colorimetric assay. 2. Begin by creating your standard curve by performing a serial dilution according to the chart below. The Insulin standard is supplied at a concentration of 300 mg/mL. Prepare spec tube #1 as indicated in the table. Add 3 mL of H2O to the remaining 4 spec tubes. Calculate what volume of Tube #1 needs to be transferred to Tube #2 etc. Table1 1 A B C D Tube mL H2O mL Standard or tube #x Concentration (mg/mL) 2 1 0 6ml of standard 300 3 2 3 3ml of Tube 1 150 4 3 3 3ml of tube 2 75 5 4 3 3ml of tube 3 37.5 6 5 3 3ml of tube 4 18.75 On this page: Entry Page 24 1ofof344 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:39 UTC 05 Jan 2025 20:24:54 UTC Table2 TUBE Concentratio n SPEC result 1 Tube 1 300 1.2 2 Tube 2 150 0.5 3 Tube 3 75 0.2 4 Tube 4 37.5 0.1 5 Tube 5 18.75 0.05 6 Unknown 1 0.1 7 Unknown 2 0.4 output (1).png Using the following equation it was possible to find unknown concentrations Concentration U1 0.1=0.0041x−0.0681 x=40.24 Concentration U2 0.4=0.0041x−0.0681 x=110.98 1. Tube #5 will be the only tube with 6 mL (the rest have 3 mL). Remove 3 mL from tube #5 and discard in the waste container. a. Add 3 mL of each spent media (both experimental and control) to separate spec tubes. b. Add 3 mL of water to a spec tube to act as a blank c. You should now have 8 spec tubes d. Add 70 µL of colour reagent (labelled “CR”) to each spec tube and swirl gently to mix On this page: Entry Page 25 2ofof344 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:39 UTC 05 Jan 2025 20:24:54 UTC e. Blank the Spec 20 by performing the following: I. With the sample holder empty, turn the left knob until the machine is "zeroed" (i.e. the needle lines up with 0% transmitted) II. Wipe the Blank with kimwipe and insert into sample holder, closing the lid III. Using the right knob, adjust the needle to 100% transmitted f. Now measure each one of your serial dilution tubes, wiping the tube with a kimwipe before inserting in sample holder. Record the absorbance of each tube. g. Measure the two spent media sample tubes in the same manner and record the absorbance. Microplate Reader: a. Using the spent media samples and the standards you prepared for the Spec 20, load 100 μL of each sample (including the blank) in triplicate in the microplate. Load the plate as indicated below. The plate will be read on a microplate reader and the values provided to you on OWL. Fill in your name(s) on the microplate template sheet and make note in your records which plate # you loaded your samples on. b. When you are done, place the spec tubes in the bin in the sink (carefully as they are expensive!) and the serological pipettes in the washer, tips up at the front of the room. Leave all the reagents in your tube rack. c. Results will be posted on OWL after lab. You will need to enter your concentration of insulin in the spent media samples (both control and experimental) on a Google sheets to collect class data. More info will come in an announcement. On this page: Entry Page 26 3ofof344 Page Entry ID null Last Modified On Export Generated on 04 Nov 2024 00:48:39 UTC 05 Jan 2025 20:24:54 UTC image.png On this page: Entry Page 27 4ofof344 Page Entry ID null Last Modified On Export Generated on 02 Nov 2024 02:11:22 UTC 05 Jan 2025 20:24:54 UTC Determining Insert Orientation in pAMP Plasmid Using Restriction Enzyme Mapping and Gel Electrophoresis Project: Molecular biology lab book Author: Kyuwon Jung Entry Created On: 02 Nov 2024 02:09:15 UTC Entry Last Modified: 02 Nov 2024 02:10:11 UTC Export Generated On: 05 Jan 2025 20:24:54 UTC FRIDAY, 2024-11-01 Restriction Digest Kit name: FastDigest, From: Thermofisher 1. Work in partners.As a pair you will complete the following reactions 2. Get four 1.5 mL microcentrifuge tubes and label them “Uncut”, “HindIII ”, “BamHI”, “and “H +B” and your initials 3. Look at the four tables below and add the appropriate components to each tube: Table1 A B 1 Component: Uncut Volume 2 Autoclaved dH2O 6 uL 3 5x FastDigest Buffer with Loading Dye 2 uL 4 250 ng Plasmid DNA. 2 uL TOTAL 10 uL 5 6 On this page: Entry Page 28 1ofof347 Page Entry ID null Last Modified On Export Generated on 02 Nov 2024 02:11:22 UTC 05 Jan 2025 20:24:54 UTC Table2 A B 1 Component: HindIII Volume 2 Autoclaved dH2O 4 uL 3 5x FastDigest Buffer with Loading Dye 2 uL 4 250 ng Plasmid DNA. 2 uL 5 HindIII 2 uL 6 TOTAL 10 uL Table3 A B 1 Component: BamHI Volume 2 Autoclaved dH2O 4 uL 3 5x FastDigest Buffer with Loading Dye 2 uL 4 250 ng Plasmid DNA. 2 uL 5 BamHI 2 uL 6 TOTAL 10 uL On this page: Entry Page 29 2ofof347 Page Entry ID null Last Modified On Export Generated on 02 Nov 2024 02:11:22 UTC 05 Jan 2025 20:24:54 UTC Table4 A B 1 Component: H +B Volume 2 Autoclaved dH2O 2 uL 3 5x FastDigest Buffer with Loading Dye 2 uL 4 250 ng Plasmid DNA. 2 uL 5 HindIII 2 uL 6 BamHI 2 uL 7 TOTAL 10 uL 1. Gently mix the tube by pipetting up and down. If there is liquid on the sides of the tube give it a quick spin in the centrifuge. 2. Place all the digest tubes in 37 °C water bath for 15 min. Loading Gel: 1. 0.75% agarose gels containing 1X Red Safe (FroggaBio) will be already prepared and in TBE running buffer. 2. Your TA will load a 1Kb plus ladder (FroggaBio) in the first lane of the gel. Make a note of which Tank # and lane # you are loading your samples. 3. You will load 8 μL of each of your restriction digests. 4. Once all samples are loaded the gel will be run at 120V for approximately 45 min. 5. The gel picture will be posted to OWL. Expected results 1. Uncut: expected to show a single band represent the plasmid in various forms such as supercoiled, nicked or concentrated 2. HindIII+BamHI(Single Digests): Each digest expected to produce single band if the enzyme cut the plasmid once 3. HindIII+BamHI(double Digest): If the insert is in the 3`to 5` orientation, the digest should yield fragments approximately 4.5kb and 1.1 kb. Additionally, if the insert is inverse, 5` to 3` orientation, the digest yield fragments of about 3.7 to 2 kb. Results 1. Uncut: The uncut lane displays multiple bands, with the most noticeable band around the ~5,700 bp mark, matching the expected plasmid size. Multiple forms are found, most likely due to the plasmid's supercoiled, nicked, and potentially concatemeric forms. 2. HindIII Digest: This lane shows a single band of approximately 5,700 bp, indicating a single HindIII site in the plasmid. This was the predicted result for a plasmid with only one HindIII cut site. 3. BamHI Digest: A single band about 5,700 bp, similar to the HindIII digest, indicates only one BamHI site.This result is also consistent with the expected effect of a single BamHI cut site in the plasmid. 4. HidIII+BamHI (Double digest) : The double digest lane features two separate bands. On this page: Entry Page 30 3ofof347 Page Entry ID null Last Modified On Export Generated on 02 Nov 2024 02:11:22 UTC 05 Jan 2025 20:24:54 UTC Based on the ladder comparison, the bands are predicted to be 4.5 kb and 1.1 kb in size. This banding pattern suggests that the insert is oriented 3' to 5', as expected. Table5 A B C 1 Lane Observed band(bp) Expected Bands(bp) 2 Uncut ~5700(multiple form) ~5700(varied forms) 3 HindIII ~5700 ~5700 4 BamHI ~5700 ~5700 5 HindIII+BamHI ~4500, ~1100 4500,1,100 On this page: Entry Page 31 4ofof347 Page Entry ID null Last Modified On Export Generated on 02 Nov 2024 02:11:22 UTC 05 Jan 2025 20:24:54 UTC 스크린샷 2024-11-01 213028.png On this page: Entry Page 32 5ofof347 Page Entry ID null Last Modified On Export Generated on 02 Nov 2024 02:11:22 UTC 05 Jan 2025 20:24:54 UTC On this page: Entry Page 33 6ofof347 Page Entry ID null Last Modified On Export Generated on 02 Nov 2024 02:11:22 UTC 05 Jan 2025 20:24:54 UTC On this page: Entry Page 34 7ofof347 Page