Experiments

Objective:

To prepare a standard solution of Triclocarban (TCC) using water as the solvent.

Materials:

• 1.0 mg of Triclocarban(TCC)
• 10 ml of distilled water
• Test tube(27ml)
• Vortex mixer
• Water bath
• Thermometer

Procedure:

Preparation of TCC Solution

1. Measure 1.0 mg of TCC using a precision balance and record the exact mass.
2. Carefully transfer the measured TCC into a clean and dry test tube.

Dissolution of TCC

1. Add 10 ml of water to the test tube containing TCC.
2. Ensure that the water completely covers the TCC powder.
3. Attach a cap or stopper to the test tube securely.
4. Gently vortex the mixture for dissolution at 3000rpm. Ensure that the TCC powder is thoroughly dispersed in the water.

Heating the Solution

1. Set up a water bath and bring it to a temperature of 60 °C.
2. Carefully place the capped test tube containing the TCC and water mixture into the heated environment.
3. Allow the solution to heat at 60 °C for 30 minutes. (Note: Record the exact heating time for consistency.)

Observation

1. After heating, carefully remove the test tube from the water bath using appropriate tools (e.g., tongs or gloves) to prevent burns.
2. Observe the solution in the test tube.
3. Record your observation. If any turbidity is present, TCC has not dissolved. If you have a clear solution, then TCC has been dissolved.

Comments:

After adding the TCC in water, we observed that TCC did not dissolve at all. We tried heating it and vortexing after with no success. This is why we decided to change solvents.

Objectives:

To prepare a series of TCC standard solutions in acetone, perform parallel dilutions, detect absorbance peaks using a UV spectrophotometer, and establish a wavelength for absorbance measurement.

Materials:

• 1.0 mg of TCC
• 10 ml of acetone
• Graduated cylinders or pipettes for dilution
• Vortex mixer
• UV spectrophotometer
• Cuvettes (3.5 ml)
• Wavelength selection guide
• Spectrophotometer software

Procedure:

1. Preparation of Initial TCC Solution

   1. Measure 1.0 mg of TCC using a precision balance and record the exact mass.
   2. Carefully transfer the measured TCC into a clean and dry container (e.g., beaker or vial).
   3. Add 10 ml of acetonitrile to the container with TCC, creating a 100 ppm (parts per million) solution.

2. Parallel Dilution

   1. Prepare a series of dilutions of (1ppm, 5ppm, 10ppm, 15ppm, 20ppm, 30ppm) by taking an aliquot of the 100 ppm TCC solution and diluting it with acetone.
   2. To prepare the standard solutions, refer to the table below

      TCC Concentration (ppm)	Volume of 100 ppm solution (ml)	Volume of Acetonitrile (ml)
      1	0.1	9.9
      5	0.5	9.5
      10	1.0	9.0
      15	1.5	8.5
      20	2.0	8.0
      30	3.0	7.0

   3. Continue parallel dilutions until you obtain a 1 ppm TCC solution. Record the volumes and concentrations of each dilution.

3. Homogenization and Initial Measurements

   1. Homogenize each TCC solution by gently mixing with a vortex mixer at 3000rpm.
   2. Using a UV spectrophotometer, run a scan program from 210 to 300 nm to find the peak of your TCC. Take the average peak value and set it as your fixed wavelength.(We have used 260)
   3. Document the results, including the absorbance and peak information.

4. Parallel Dilution and Absorbance Measurements (25 ppb-250 ppb Range)

   1. Perform additional parallel dilutions of the TCC solution to create concentrations from 25 ppb to 250 ppb.
   2. Repeat the absorbance measurements at 260 nm for each dilution using the UV spectrophotometer. Record the absorbance and peak information.
   3. Document the results, including each concentration's absorbance and peak information within the specified range.

5. Multiple Readings and Peak Detection at 260 nm

   1. Using the fixed wavelength of 260 nm, perform multiple readings at different concentrations of TCC.
   2. Record the absorbance values for each concentration and observe the peak characteristics.
   3. Document the results, including the absorbance and peak information at 260 nm.

Comments:

The results we got were mostly negative. We suspect the reasons to be the high volatility of acetone and its similar peak wavelength (280nm). The volatility keeps changing the properties of the solution.

Objectives:

1. To prepare a series of TCC standard solutions in acetonitrile, including a 100 ppm stock solution.
2. To create standard solutions with different concentrations for calibration.

Materials:

• 1.0 mg of TCC
• 10 ml of acetonitrile
• Graduated cylinders or pipettes for dilution
• UV spectrophotometer
• Quartz cuvettes(3.5ml)
• Wavelength selection guide
• Spectrophotometer software

Procedure:

1. Preparation of Initial TCC Solution

   1. Measure 1.0 mg of TCC using a precision balance and record the exact mass.
   2. Carefully transfer the measured TCC into a clean, dry container (e.g., beaker or vial).
   3. Add 10 ml of acetonitrile to the container with TCC, creating a 100 ppm (parts per million) solution.
   4. Label the solution with the concentration (100 ppm).

2. Acetonitrile as Blank:

   Set aside a sample of acetonitrile to be used as a blank for this experiment.

3. Preparation of Standards

   Prepare a series of dilutions using the 100 ppm TCC solution and acetonitrile to achieve the desired concentrations:

   TCC Concentration (ppb)	Volume of 100 ppm solution (ml)	Volume of Acetonitrile (ml)
   50	0.005	9.995
   100	0.01	9.99
   250	0.025	9.975
   1000	0.1	9.9
   2500	0.25	9.25
   5000	0.5	9.5
   7500	0.75	9.25
   10000	0.1	9.0

4. Calculation of Limit of Detection (LOD)

   1. To determine the LOD, quality curve is required. Perform absorbance measurements for each of the prepared standards at a fixed wavelength.
   2. Record the absorbance values for each standard solution.
   3. Perform absorbance measurements for at least six different concentrations to create a calibration curve.

5. Spectrometer's Error Value

   1. Since the first absorbance value is 0.011, it is necessary to determine the spectrometer's error value. This can be done by measuring the absorbance of a blank solution (acetonitrile) and calculating the error.
   2. Record the absorbance value obtained for the blank solution.

Objective:

To revive a culture stored in a glass ampoule by transferring it into suitable growth media, distributing it into multiple containers, and incubating it under optimal conditions for further use.

Materials:

• Glass ampoule containing the culture
• Sterile blade
• Heat sterilised forceps
• 0.5ml liquid media
• Pipette
• Test tubes(27ml)
• Petri dishes with nutrient broth agar
• Incubator
• Labels

Procedure:

1. Aseptic Handling

   1. Begin by ensuring aseptic conditions in the laboratory. Disinfect the work area by wiping the surfaces with 70% ethanol, follow LAF sterilisation procedures and wash your hands thoroughly.
   2. Wear appropriate personal protective equipment (PPE), including lab coats, gloves, and safety goggles.

2. Ampoule Disintegration

   1. Take the glass ampoule containing the culture.
   2. Using a sterile blade, carefully file the neck of the ampoule to create a score mark.
   3. Hold the ampoule firmly, and using the sterile blade, break the neck along the score mark to release the culture into the bottom of the ampoule.

3. Cotton Plug Removal:

   Using heat-sterilized forceps, carefully remove the cotton plug from the ampoule.

4. Resuspension in Liquid Media

   1. Using a pipette, add 0.5 ml of the appropriate liquid growth media into the ampoule.
   2. Gently resuspend the culture by pipetting up and down, ensuring even distribution of the culture in the liquid media.

5. Distribution into Containers

   1. Distribute the revived culture into multiple containers for further growth and experimentation:
      1. Transfer a portion of the resuspended culture into two separate test tubes containing nutrient broth.
      2. Transfer another portion into two Petri dishes containing nutrient broth agar.
   2. Label each container appropriately with the culture name, date, and any other necessary information.

6. Incubation

   Place the labelled containers into an incubator set to the required optimum temperature and time period as specified by the culture suppliers.

7. Monitoring

   1. Periodically monitor the containers for growth and observe for any signs of contamination.
   2. Note any changes in appearance, such as the presence of colonies or turbidity in the liquid media.

8. Streaking of a new plate

   1. Take a heat sterilized inoculation loop and dip it into the eppendorf tube containing the culture.
   2. Quadrant streak the culture present on the loop onto a petri plate.

Comments:

Make sure to keep extra plates, because the pour plates we incubated had lawns of the culture, keep the extra plate to streak it as well or to add less concentration of the pure culture.

Objective:

To perform the catalase test and determine if the colony exhibits a positive catalase reaction.

Materials:

• Cell colony
• Clean, dry glass slide
• Loop or sterile wooden stick
• 3% hydrogen peroxide (H2O2)
• Biohazard glass disposal container

Procedure:

1. Preparation

   1. Begin by ensuring proper laboratory safety measures. Wear appropriate personal protective equipment (PPE), including lab coats and safety goggles.
   2. Work in a designated area for microbiological tests.

2. Colony Transfer

   Using a loop or a sterile wooden stick, transfer a small amount of the colony to the surface of a clean, dry glass slide.

3. Catalase Reaction Test

   1. Place a drop of 3% hydrogen peroxide (H2O2) directly onto the colony on the glass slide.
   2. Mix the colony and hydrogen peroxide gently by gently stirring or tapping with the loop or sterile stick.
   3. Observe the reaction carefully
   4. Look for the rapid evolution of oxygen, which is evidenced by the formation of bubbles. This should occur within 5-10 seconds.
   5. If bubbling occurs within the specified time frame, the test is considered positive for catalase activity.

4. Disposal:

   Dispose of the glass slide used for the test in a designated biohazard glass disposal container.

Objective:

To perform a Gram stain on a bacterial smear and observe the results, specifically identifying the bacteria as Gram-negative.

Materials:

• Clean glass slide
• Bacterial culture or smear
• Bunsen burner or slide warmer
• Crystal violet stain
• Iodine solution
• 95% ethyl alcohol (decolorization solution)
• Safranin stain
• Microscope with oil immersion objective lens
• Microscope immersion oil
• Distilled water or tap water

Procedure:

1. Preperation of Smear

   1. Take a clean glass slide and place a small amount of the bacterial culture or smear on it.
   2. Using a sterile loop or wire, spread the bacterial material into a thin, even smear on the slide.
   3. Allow the smear to air dry completely.
   4. Heat fix the smear by passing it through the flame of a Bunsen burner or using a slide warmer. Ensure that the slide is not overheated, which could lead to distortion of bacterial morphology.

2. Staining Procedure

   1. Flood the heat-fixed smear with crystal violet stain, ensuring that the entire smear is covered. Allow the crystal violet to stand for 1 minute.
   2. Rinse the slide with tap water to remove excess crystal violet.
   3. Flood the smear with iodine solution, covering the entire smear. Allow the iodine to stand for 1 minute. The iodine serves as a mordant and enhances staining.
   4. Rinse the slide with tap water to remove excess iodine.
   5. Decolorize the smear by gently flooding it with 95% ethyl alcohol. Allow the alcohol to stand for approximately 10 seconds. Monitor the decolorization closely, as over-decolorization can lead to false results.
   6. Rinse the slide with tap water immediately to stop the decolorization process.
   7. Counterstain the smear by flooding it with safranin stain. Allow the safranin to stand for 45 seconds.
   8. Rinse the slide with tap water to remove excess safranin.

3. Observation

   1. Allow the slide to air dry completely.
   2. Examine the dried slide under a microscope using the oil immersion objective lens.
   3. Observe the bacterial morphology and the color of the cells after staining.

4. Gram Staining Result

   Based on the observation, record the Gram staining result. In this case, if the cells appear pink or red, record the result as Gram-negative.

Objective:

To perform the indole spot test on a culture and determine the result as negative for the presence of indole.

Materials:

• Cell culture (18-24 hour growth)
• Petri dish cover
• Whatman filter paper
• Kovács reagent

Procedure:

1. Preparation of cell culture:

   Inoculate a culture and incubate it for 18 to 24 hours at 31 °C to allow for growth.

2. Indole Spot Test Preparation

   1. To conduct the indole spot test, begin by placing a small piece of Whatman filter paper in a clean and dry petri dish cover.
   2. Saturate the filter paper with Kovács reagent, adding approximately 1 to 1.5 ml of the reagent to ensure the paper is thoroughly soaked.

3. Smeared Sample Preparation

   1. Obtain a small amount of cell paste from the 18 to 24-hour culture. This can be achieved by gently scraping a loop or sterile wooden stick along the surface of the culture.
   2. Smear the cell paste onto the saturated Whatman filter paper in the petri dish cover.

4. Incubation

   1. Carefully cover the petri dish to create a closed environment.
   2. Incubate the prepared filter paper at the appropriate temperature and time period (31 °C for 18-24 hours).

5. Observation:

   After the incubation period, carefully examine the filter paper for any color change or visible indication of indole production.

6. Indole Spot Test Result:

   Based on the observation, record the result of the indole spot test. If there is no visible color change or indication of indole production, record the result as negative.

Objective:

To prepare Methyl Red and Voges-Proskauer (MR-VP) broth medium, inoculate it and incubate the test tubes at 37 °C for 24 hours in preparation for the Methyl Red and Voges-Proskauer tests.

Materials:

• MR-VP broth medium (prepared in advance)
• Test tubes (three)(27 ml)
• Autoclave
• Culture
• Inoculation loop or sterile wooden stick
• Static incubator set at 37 °C
• Labels

Procedure:

1. Preparation of MR-VP Broth Medium

   1. Prepare MR-VP broth medium in three separate test tubes.
   2. Ensure that the medium is properly mixed and sterilized by autoclaving.
   3. Label the test tubes as "MR," "VP," and "Control" for identification.

2. Inoculation of Culture

   1. Using sterile technique, ensure that all equipment and materials are free from contamination.
   2. Inoculate each labeled test tube with bacterial culture as follows:
      1. Using an inoculation loop or sterile wooden stick, transfer a small amount of bacterial culture into the "MR" labeled test tube containing MR-VP broth medium.
      2. Similarly, inoculate the "VP" labeled test tube with bacterial culture.
      3. The "Control" tube will not receive any inoculation and will serve as a control.

3. Incubation

   1. Place all the labeled test tubes in an incubator set at 37 °C.
   2. Allow the test tubes to incubate for a period of 24 hours.

4. Monitoring

   1. Periodically check the test tubes during the incubation period to ensure that they remain at the correct temperature.
   2. Observe for any signs of growth or changes in the medium.

Objective:

To prepare a series of TCC standard solutions using a mixture of 99% water and 1% TCC in acetonitrile, perform dilutions, and detect the absorbance of these samples at 260nm using a UV spectrophotometer.

Materials:

• 1.0mg of TCC
• 10ml acetonitrile
• 10ml water
• Graduated cylinders or pipettes for dilution
• UV spectrophotometer
• Cuvettes (3.5 ml)
• Wavelength selection guide
• Spectrophotometer software
• Labels

Procedure:

1. Preparation of Initial TCC Solution

   1. Measure 1.0 mg of TCC using a precision balance and record the exact mass.
   2. Carefully transfer the measured TCC into a clean and dry container (e.g., beaker or vial).
   3. Add 10 ml of acetonitrile to the container with TCC, creating a 100 ppm (parts per million) solution.

2. Blank Solution:

   Prepare a blank solution using acetonitrile, which will serve as a reference during absorbance measurements.

3. Dilution for 10 ppm Solution

   1. Pipette 0.1 ml of the 100 ppm TCC solution into a clean test tube.
   2. Add 9 ml of the blank solution to the same test tube, creating a 10ppm TCC solution.

4. Preparation of 100 ppb-3000 ppb Samples

   1. Pipette 0.1 ml of the 10 ppm TCC solution into a clean test tube containing 9.9 ml of water. This creates a 100 ppb TCC solution.
   2. Repeat the process, each time using the 10 ppm TCC solution but varying the volume of water added as follows:
      1. 0.25 ml of solution 2 with 9.75 ml of water (250 ppb).
      2. 0.5 ml of solution 2 with 9.5 ml of water (500 ppb).
      3. 1 ml of solution 2 with 9 ml of water (1000 ppb or 1 ppm).
      4. 2 ml of solution 2 with 8 ml of water (2000 ppb or 2 ppm).
      5. 3 ml of solution 2 with 7 ml of water (3000 ppb or 3 ppm).

5. Absorbance Measurements

   1. Using a UV spectrophotometer, set the wavelength to 260 nm.
   2. Prepare cuvettes with the various TCC standard solutions and the blank solution.
   3. Measure the absorbance of each sample at 260 nm using the UV spectrophotometer.

Objective:

To create a growth curve for bacteria in the presence of acetonitrile by measuring optical density (OD) at different time intervals.

Materials:

• Three conical flasks
• 25 ml LB broth per flask
• Autoclave
• 1 ml overnight culture of bacterial
• 1% acetonitrile solution
• Spectrophotometer
• Cuvettes (3.5 ml)
• Timer or stopwatch
• Labels

Procedure:

1. Preparation of LB Broth

   1. Take three conical flasks and add 25 ml of LB broth to each flask.
   2. Autoclave the flasks to sterilize the LB broth medium.

2. Flask Setup

   1. Label the three flasks as follows: "Blank," "Control," and "Culture + Acetonitrile."
   2. Prepare the contents of each flask as follows:
      1. Blank: Contains only LB broth medium.
      2. Control: Add 1 ml of the overnight culture of bacteria to the LB broth. Mix well.
      3. Culture + Acetonitrile: Add 1 ml of the culture to the LB broth and then add 1% acetonitrile solution to the same flask. Mix well.

3. Incubation

   1. Place each flask in an incubator set to the appropriate temperature for the growth of bacteria.
   2. incubate the flasks under controlled conditions.

4. Observation and OD Measurements

   1. At regular intervals (e.g., every hour), remove each flask from the incubator.
   2. Using a spectrophotometer, measure the optical density (OD) of the cultures at 600nm.
   3. Record the OD readings for each flask at each time interval.
   4. Keep the cultures in the incubator between measurements.

5. Growth Curve Plotting:

   Plot the OD readings against time to create a growth curve for bacteria in the presence of acetonitrile.

6. Data Analysis

   Analyze the growth curve to observe and interpret the growth pattern and the effect of acetonitrile on bacterial growth.

Objective:

To prepare a series of TCC standard solutions for LCMS (Liquid Chromatography-Mass Spectrometry) analysis by diluting a stock TCC solution in acetonitrile, and then mixing it with water to achieve various concentrations ranging from 100 ppm to 3000 ppb

Materials:

• 1.0 mg of TCC
• 10 ml acetonitrile
• 9 ml water
• Pipettes (0.1 ml, 0.05 ml)
• Test tubes(27ml)
• Labels

Procedure:

1. Preparation of Stock TCC Solution

   1. Measure precisely 1.0 mg of TCC using a precision balance and record the exact mass.
   2. Transfer the measured TCC into a test tube.
   3. Add 10 ml of acetonitrile to the container with TCC, creating a 100 ppm (parts per million) TCC stock solution. Label the solution with its concentration (100 ppm).

2. Blank Solution:

   Prepare a blank solution using 10 ml of acetonitrile. This solution will serve as a reference during LCMS analysis.

3. Preparation of 500 ppb-5000 ppb Samples

   1. Pipette 0.05 ml of Solution into a clean test tube labeled "Solution 1."
   2. Pipette 0.1 ml of Solution into a clean test tube labeled "Solution 2."
   3. Pipette 0.2 ml of Solution into a clean test tube labeled "Solution 3."
   4. Pipette 0.3 ml of Solution into a clean test tube labeled "Solution 4."
   5. Pipette 0.4 ml of Solution into a clean test tube labeled "Solution 5."
   6. Pipette 0.5 ml of Solution into a clean test tube labeled "Solution 6."

4. Final Dilution

   To each test tube containing the various solutions (1 to 6), add enough water to reach a total volume of 10 ml, ensuring proper mixing.

Objective:

To create a growth curve for bacterial culture at a chosen pH by measuring optical density (OD) at different time intervals.

Materials:

• Four conical flasks
• 50 ml LB broth per flask
• Autoclave
• 1N NaOH solution
• 1N HCl
• Bacterial culture culture
• Spectrophotometer
• Cuvettes (3.5 ml)
• Timer or stopwatch
• pH meter
• Labels

Procedure:

1. Preparation of LB Broth at chosen pH

   1. Take four conical flasks and add 50 ml of LB broth to each flask
   2. Adjust the pH of the LB broth in the flasks to chosen pH using 1N NaOH or 1N HCl respectively.
   3. Autoclave the flasks to sterilize the LB broth medium.

2. Flask Setup

   1. Label the four flasks as follows: "Blank," "chosen pH - 1," "chosen pH - 2," and "chosen pH - 3."
   2. Prepare the contents of each flask as follows:
      1. Blank: Contains only LB broth medium at chosen pH.
      2. Chosen pH- 1: Inoculate the flask with one colony of bacterial culture. Mix well.
      3. Chosen pH- 2: Inoculate the flask with another colony of bacterial culture. Mix well.
      4. Chosen pH - 3: Inoculate the flask with a third colony of bacterial culture. Mix well.

3. Incubation

   1. Place each flask in an incubator set to the appropriate temperature for the growth of bacteria.
   2. Incubate the flasks under controlled conditions.

4. Observation and OD Measurements

   1. At regular intervals (e.g., every hour), remove each flask from the incubator.
   2. Measure the optical density (OD) of the cultures at 600nm using a spectrophotometer.
   3. Record the OD readings for each flask at each time interval.
   4. Keep the cultures in the incubator between measurements.

5. Growth Curve Plotting:

   Plot the OD readings against time to create a growth curve for bacteria at chosen pH.

6. Data Analysis

   Analyze the growth curve to observe and interpret the growth pattern of bacteria at chosen pH.

Objective:

To dissolve a lyophilized IDT tccA gene fragment, measure its concentration, and prepare a stock solution for storage at -20°C.

Materials:

• Lyophilized IDT tccA gene fragment
• 100 μl autoclaved Milli-Q water
• Microcentrifuge tube
• Vortex mixer
• Centrifuge
• Ice
• UV-Vis spectrophotometer
• Cuvette(3.5 ml)
• Pipettes (25 μl, 75 μl)
• Milli-Q water
• -20°C freezer
• Labels

Procedure:

1. Dissolving the Lyophilized Gene Fragment

   1. Add 100 μl of autoclaved Milli-Q water to the lyophilized IDT tccA gene fragment in a microcentrifuge tube.
   2. Very gently shake the tube to ensure the gene fragment comes into contact with the water.
   3. Vortex the sample for 1 second to aid in dissolving the gene fragment.
   4. Centrifuge the sample at 14,000 rpm for 30 seconds to pellet any remaining undissolved material.
   5. Immediately place the sample in an ice bath to maintain its integrity.

2. Measurement of OD (Optical Density):

   Measure the optical density (OD) of the sample using a UV-Vis spectrophotometer at 260nm for nucleic acids.

3. Dilution of Gene Fragment:

   1. Pipette 25 μl of the dissolved gene fragment solution and transfer it into a new microcentrifuge tube.
   2. Add 75 μl of Milli-Q water to the tube, resulting in a total volume of 100 μl.
   3. Ensure that the diluted solution is thoroughly mixed.

4. Concentration Calculation:

   Calculate the concentration of the dissolved gene fragment. For example, if the OD measurement indicates a concentration of 10 ng/μl, record this value.

5. Storage

   1. Label the microcentrifuge tube containing the dissolved gene fragment with the concentration value (e.g., 10 ng/μl) and any other relevant information.
   2. Store the tube in a -20°C freezer for long-term storage.

Objective:

To dissolve lyophilized tccA primers, measure their concentration, and prepare a stock solution for storage at -20°C.

Materials:

• Lyophilized Forward Primer
• Lyophilized tccA Reverse Primer
• Autoclaved Milli-Q water
• Microcentrifuge tubes (two)
• Vortex mixer
• Centrifuge
• Ice
• UV-Vis spectrophotometer
• Cuvette (3.5 ml)
• Pipettes (10 μl, 90 μl)
• Milli-Q water
• -20°C freezer
• Labels

Procedure:

1. Dissolving the Lyophilized Primers

   1. Add 368.9 μl of autoclaved Milli-Q water to the lyophilized Forward Primer (100 pmol/ μL) in a microcentrifuge tube.
   2. Add 249.3 μl of autoclaved Milli-Q water to the lyophilized tccA Reverse Primer (100 pmol/ μL) in a separate microcentrifuge tube.
   3. Gently mix each tube to ensure the primer comes into contact with the water.
   4. Vortex each sample for 1 second to aid in dissolving the primers.
   5. Centrifuge each sample at 14,000 rpm for 30 seconds to pellet any remaining undissolved material.
   6. Immediately place each sample in an ice bath to maintain primer integrity.

2. Dilution of Primers

   1. For each primer, pipette 10 μl of the dissolved primer solution and transfer it into a new microcentrifuge tube.
   2. Add 90 μl of Milli-Q water to each tube, resulting in a total volume of 100 μl.
   3. Gently pipette mix each tube to ensure thorough mixing.

3. Measurement of Concentration

   1. Measure the concentration of each primer using a UV-Vis spectrophotometer at an appropriate wavelength for nucleic acids (e.g., 260 nm).
   2. Verify that the concentration of each primer is approximately 10 μM.

4. Storage

   1. Label each microcentrifuge tube containing the dissolved primer with the primer name and a concentration value (e.g., Forward Primer 10 μM, tccA Reverse Primer 10 μM).
   2. Store the tubes in a -20°C freezer for long-term storage.

Objective:

To prepare PCR reaction mixtures for tccA gene amplification, including master mix preparation, dividing it into separate tubes, and creating samples for both negative and positive controls.

Materials:

• PCR reagents (e.g., Taq polymerase, dNTPs, buffer, primers)
• Template DNA
• Microcentrifuge tubes (0.2 ml)
• Pipettes (various sizes)
• PCR tube rack
• Labels

Procedure:

1. Master Mix Preparation

   1. Prepare a master mix for a 50 μl PCR reaction mixture in a 0.2 ml PCR tube. The master mix should include all necessary PCR reagents, excluding the template DNA and primers as mentioned below.
   2. Mix the components of the master mix thoroughly to ensure homogeneity.
   3. Calculate the total volume of the master mix, which should be 50 μl.

2. Dividing the Master Mix:

   Divide the total 50 μl PCR reaction mixture into two separate 0.2 ml PCR tubes, with each tube containing 25 μl of the master mix.

3. Sample Creation:

   Divide each of the two 25 μl PCR reaction mixtures into three separate samples. You will have a total of six samples.

4. Control Samples

   1. For the negative control samples:
      1. In one of the 25 μl reaction mixtures, add no template DNA. This will serve as the "No template DNA" negative control.
      2. In another 25 μl reaction mixture, add all components of the master mix except the DNA polymerase (e.g., Q5 High-Fidelity DNA Polymerase). This will serve as the "No Q5" negative control.
   2. In another 25 μl reaction mixture, add all components of the master mix except the DNA polymerase (e.g., Q5 High-Fidelity DNA Polymerase). This will serve as the "No Q5" negative control.

5. Labeling:

   Label each sample tube appropriately, indicating the type of sample (e.g., Negative Control - No template DNA, Negative Control - No Q5, Positive Control - Mam7 gene) and any other relevant information.

Objective:

To prepare a master mix for gradient PCR, aliquot it into PCR tubes for multiple reactions, and set up samples for gradient PCR, including negative and positive controls.

Materials:

• 2X Q5 High-Fidelity Master Mix
• PCR primers
• Template DNA (if required)
• Autoclaved Milli-Q water
• Eppendorf tube
• PCR tubes (9 or as required)
• Pipettes (various sizes)
• PCR tube rack
• Labels

Procedure

1. Master Mix Calculation:

   Calculate the amount of each component required for the master mix, including water, 2X Q5 High-Fidelity Master Mix, and primers. Ensure that the master mix is sufficient for n+1 reactions, where n is the number of reactions required.

2. Master Mix Preparation

   1. Prepare the master mix by combining the calculated amounts of water, 2X Q5 High-Fidelity Master Mix, and primers in an Eppendorf tube.
   2. Mix the components thoroughly to ensure homogeneity.

3. Aliquoting the Master Mix:

   Aliquot the prepared master mix into 9 separate PCR tubes. These tubes will be used for 8 samples for gradient PCR and one for a negative control (sample 11) with no DNA.

4. Sample Preparation:

   Set up the following samples in the PCR tubes containing the master mix:

   • Samples 1 to 8: These are your experimental samples for gradient PCR. Ensure that each tube contains a different DNA template concentration, as per your gradient PCR design.
   • Sample 9: Negative control with 10 ng of tccA (if applicable) and water. Mix thoroughly.
   • Sample 10: Positive control with the MAM7 gene (if applicable). Mix thoroughly.
   • Sample 11: Negative control with no DNA. This serves as a blank control.

5. Labeling:

   Label each PCR tube appropriately, indicating the sample number, type of sample, and any other relevant information.

Objective:

To isolate plasmid DNA from bacterial cells using the QIAprep 2.0 spin column-based plasmid isolation method and measure the concentration of the isolated plasmid.

Materials:

• Bacterial culture samples (1-10)
• QIAprep 2.0 spin column kit
• Microcentrifuge tubes
• Pipettes (various sizes)
• Table-top microcentrifuge
• Collection tubes
• Buffer P1
• Buffer P2
• Buffer N3
• Buffer PE
• Buffer EB (10 mM Tris-HCl, pH 8.5)
• Spectrophotometer
• Labels

Procedure:

1. Pelleting Bacterial Cells:

   Centrifuge the bacterial culture samples (1-10) at 13,000 rpm for 10 minutes at 4°C to pellet the bacterial cells.

2. Resuspension and Lysis

   1. Resuspend the pelleted bacterial cells in 250 μl of Buffer P1. Transfer the resuspended cells to a microcentrifuge tube.
   2. Add 250 μl of Buffer P2 to the cell suspension. Mix thoroughly by inverting the tube 4–6 times until the solution becomes clear.
   3. Add 350 μl of Buffer N3 immediately and mix immediately and thoroughly by inverting the tube 4–6 times.
   4. Centrifuge the mixture for 10 minutes at 13,000 rpm (~17,900 x g) in a table-top microcentrifuge.

3. Binding and Washing

   1. Add 800 μl of the supernatant to a QIAprep 2.0 spin column by pipetting.
   2. Centrifuge the column for 30–60 seconds at 13,000 rpm and discard the flow-through.
   3. Wash the QIAprep 2.0 spin column by adding 0.75 ml of Buffer PE. Centrifuge for 30–60 seconds at 13,000 rpm.

4. Elution of Plasmid DNA

   1. Transfer the QIAprep 2.0 spin column to a collection tube.
   2. Centrifuge the column for 1 minute to remove the residual wash buffer.
   3. Place the QIAprep 2.0 column in a clean 1.5 ml microcentrifuge tube.
   4. To elute DNA, add 30 μl of Buffer EB (10 mM Tris-HCl, pH 8.5) to the center of the QIAprep 2.0 spin column. Let it stand for 5 minutes, then centrifuge for 1 minute at 13,000 rpm.
   5. Repeat the elution step by adding another 15 μl of Buffer EB to the column, letting it stand for 4 minutes, and then centrifuging for 1 minute at 13,000 rpm.

5. Measurement of Plasmid Concentration:

   Measure the concentration of the isolated plasmid using a spectrophotometer at 260nm.

Objective:

To purify amplified tccA DNA fragments using a QIAquick column-based PCR cleanup method and determine the gene's concentration by UV spectrophotometry.

Materials:

• Amplified tccA DNA
• Buffer PB
• QIAquick column
• 2 ml collection tube
• Buffer PE
• Autoclaved 1.5 ml Eppendorf tube
• Buffer EB (10 mM Tris·Cl, pH 8.5)
• UV spectrophotometer

Procedure:

1. Purification of Amplified tccA DNA

   1. Take each separate 25 μL of the amplified tccA DNA
   2. Add 125 μL of Buffer PB to each of the amplified tccA mixes and mix thoroughly.
   3. Transfer each mixture to a separate QIAquick column, placing each column in a provided 2 ml collection tube.
   4. Centrifuge each column for 30–60 seconds at 13,000 rpm, 32°C.
   5. Discard the flow-through and place the QIAquick columns back in their respective collection tubes.
   6. Add 750 μL of Buffer PE to each QIAquick column and centrifuge for 30–60 seconds at 13,000 rpm, 32°C for washing.
   7. Centrifuge each QIAquick column for an additional 1 minute at 13,000 rpm, 32°C in a new 2 ml collection tube to remove residual wash buffer.
   8. Place each QIAquick column in an autoclaved 1.5 ml Eppendorf tube.
   9. Add 25 μL of Buffer EB (10 mM Tris·Cl, pH 8.5) to the center of each QIAquick membrane and let it stand for 1 minute.
   10. Centrifuge each QIAquick column for 1 minute at 13,000 rpm, 32°C to elute the DNA.
   11. To the same QIAquick membrane, add 15 μL of Buffer EB to the center and place it in a new 2 ml collection tube that was provided. Centrifuge for 1 minute at 13,000 rpm, 32°C to elute the DNA again.

2. Concentration Determination by UV Spectrophotometry:

   Measure the concentration of the purified gene using a UV spectrophotometer at 260 nm.

Objective:

To test the effectiveness of enzyme digestion using both single and double digestion reactions.

Materials:

• Template DNA (for 250 ng of vector)
• 10X CutSmart Buffer
• BamHI enzyme
• XhoI enzyme
• Milli-Q water
• Microcentrifuge tubes
• PCR tube rack
• Thermal cycler
• DNA ladder (1 kb ladder)
• Gel electrophoresis apparatus
• Gel-loading dye

Procedure

• Single Digestion Reaction Mix Preparation

  1. Components for the single digestion of the gene:
     • Template DNA (for 250 ng insert): 10 μl
     • 10X CutSmart Buffer: 2.5 μl
     • Restriction enzyme of choice: 0.25 μl
     • Milli-Q water: Make up to 25 μl

  2. Prepare the single digestion reaction mixtures as specified above and mix them thoroughly. Incubate the reactions at the appropriate temperature for restriction enzyme digestion (typically 37°C) for 3 hours

• Double Digestion Reaction Mix Preparation

  1. Components for the double digestion of the gene and plasmid vector:
     • Template DNA (for 250 ng insert): 10 μl
     • 10X CutSmart Buffer: 2.5 μl
     • Restriction enzyme 1 of choice: 0.25 μl
     • Restriction enzyme 2 of choice: 0.25 μl
     • Milli-Q water: Make up to 25 μl
  2. Prepare the double digestion reaction mixture as specified above and mix it thoroughly.

• Thermocycling Conditions for Double Digestion

  1. Perform the following thermocycling steps for the double digestion reaction:
     • Incubation at 37°C for 3 hours for enzyme digestion.
     • Enzyme inactivation at 65°C for 20 minutes.
     • Hold the reaction at 4°C until further use.

• Gel Electrophoresis

  1. Load the digested DNA samples along with a 1 kb DNA ladder onto an agarose gel.
  2. Perform gel electrophoresis using appropriate electrophoresis conditions

Objective:

To perform Gibson assembly by combining the required components to create a reaction mixture and follow the protocol provided in the NEB (New England Biolabs) Gibson Assembly protocol (Reference: NEB Gibson Assembly Protocol).

Materials

• Gibson Assembly Master Mix (2X)
• Template DNA (50 ng/μl)
• Insert DNA (38 ng/μl)
• Nuclease-free water
• Microcentrifuge tubes
• Pipettes (10 μl, 2 μl, 0.25 μl, 7.25 μl)
• thermocycler
• Incubator (50°C)

Procedure

1. Component Preparation:

   Component	Volume(μl)	Concentration Available
   Gibson Assembly Master Mix	10	2X
   Template DNA	2	50 ng/μl
   Insert DNA	0.25	38 ng/μl
   Nuclease-free Water	7.25
   Total Volume	20

2. Reaction Mix Preparation

   1. In a microcentrifuge tube, add the following components in the order listed:
      • 10 μl of Gibson Assembly Master Mix (2X)
      • 2 μl of Template DNA
      • 0.25 μl of Insert DNA
      • 7.25 μl of Nuclease-free Water
   2. Mix the components gently by pipetting up and down, ensuring thorough mixing. Avoid introducing air bubbles.

3. Gibson Assembly Reaction

   1. Follow the Gibson Assembly protocol provided in the NEB reference link (here) for the specific temperature and incubation times required for the reaction.
   2. Typically, this will involve incubating the reaction mixture at 50°C for a certain duration to allow for assembly and then cooling to 4°C or placing on ice to terminate the reaction.

4. Transformation (If Required):

   If the next step involves transforming the assembled DNA into a host organism (e.g., E. coli), follow the appropriate transformation protocol.

Objective:

To transform Gibson assembled tccA + pET22b+ into E. coli DH5alpha using the CaCl2 method, followed by plating and incubation.

Materials:

• 0.1M CaCl2
• Anhydrous CaCl2 (11.1g)
• Autoclaved distilled water
• PVDF filter (0.22μm)
• LB Agar
• LB agar powder (6g)
• Ampicillin (0.25μL)
• Distilled water
• LB Broth
• LB broth powder (2.5g)
• E. coli DH5-alpha culture
• Glycerol stock
• Falcon tubes (15ml)
• Ice
• 1.5ml Falcon tubes
• Plasmid (10ng)
• LB media (pre-warmed)
• LB ampicillin plates

Procedure:

1. Chemical and Media Preparation

   1. 0.1M CaCl2 a. Weigh out 11.1g of anhydrous CaCl2 and add it to 80mL of autoclaved distilled water. b. Mix the solution until the CaCl2 is fully dissolved. c. Top up the solution to a final volume of 100mL. d. Take 10mL of the above solution and dilute it by adding 90mL of autoclaved distilled water for a 1:10 dilution. e. Filter sterilized the solution through a PVDF filter with a pore size of 0.22μm.
   2. LB Ampicillin Plates a. Weigh out 6g of LB agar powder and add it to 250ml of distilled water. b. Autoclave the LB agar solution. c. Dissolve 0.25μL of ampicillin into the LB agar solution. d. Pour 25ml of LB-ampicillin solution into each Petri plate.
   3. LB Ampicillin Plates a. Weigh out 6g of LB agar powder and add it to 250ml of distilled water. b. Autoclave the LB agar solution. c. Dissolve 0.25μL of ampicillin into the LB agar solution. d. Pour 25ml of LB-ampicillin solution into each Petri plate.

2. E. coli DH5-alpha Culture Preparation

   Inoculate 200μL of E. coli DH5-alpha from the glycerol stock onto 5ml of LB. b. Incubate the culture at 37°C and 200rpm for 16 hours. c. Measure the growth of the DH5-alpha culture using a spectrophotometer at absorbance A600. d. Transfer the culture to 15ml falcon tubes placed on ice. e. Centrifuge the falcon tubes at 4°C at 5600g for 10 minutes to pellet the DH5-alpha culture. f. Discard the supernatant. g. Resuspend each pellet in 1ml of ice-cold 0.1M CaCl2. h. Incubate the 2ml cell suspension on ice for 30 minutes. i. Centrifuge the suspension at 4°C at 5600g for 10 minutes. j. Discard the supernatant. k. Resuspend each pellet in 500μL of ice-cold 0.1M CaCl2. l. Prepare five 100μL aliquots of competent cells in ice-cold 1.5ml falcon tubes. m. Store the prepared competent cells at -80°C.

3. Transformation

   • Add 10ng of plasmid to each aliquot for the required transformant and positive control.
   • Incubate the cells on ice for 30 minutes.
   • Place the cells in a 42°C water bath for exactly 30 seconds.
   • Place the cells on ice for 2 minutes.
   • Add 1ml of pre-warmed LB media to each aliquot.
   • Incubate the aliquots of cells at 37°C, 200rpm for 1 hour.

4. Plating and Incubation

   • Spread plate 100μL of the transformed cells from the aliquots onto LB ampicillin plates.
   • Incubate the plates at 37°C for 12-16 hours.

Objective:

To perform colony PCR of transformed cells, including colony cell suspension preparation, PCR reaction mix preparation, and setting thermocycling conditions.

Materials:

• Transformed cell colonies
• Autoclaved toothpick
• Eppendorf tubes
• Nuclease-free (autoclaved Milli-Q) water
• PCR reagents (e.g., Q5 Master Mix, T7 Forward Primer, tccA Reverse Primer)
• PCR tubes
• Thermocycler
• Labels

Procedure:

1. Colony Cell Suspension Preparation

   1. Using a freshly autoclaved toothpick, touch it onto a colony of transformed cells.
   2. Dip the toothpick into an Eppendorf tube containing 100 μl of nuclease-free (autoclaved Milli-Q) water to create a colony cell suspension. Ensure the toothpick is fully submerged in the water.
   3. Mix the suspension by gentle vortexing or pipetting.

2. PCR Reaction Mix Preparation:

   Prepare a PCR reaction mix for colony PCR with the following components in a PCR tube:

   • Q5 Master Mix (2X): 12.5 μl
   • T7 Forward Primer: 1.5 μl
   • tccA Reverse Primer: 1.5 μl
   • Autoclaved Milli-Q Water: 8.5 μl
   • Colony Cell Suspension: 1 μl

   Mix the components by gentle pipetting or vortexing.

3. Thermocycling Conditions for Colony PCR:

   Set up the thermocycler with the following cycling conditions:

   • Initial Denaturation:
     • Temperature: 98°C
     • Time: 3 minutes
   • Cycling (30 cycles):
     • Denaturation
       • Temperature: 98°C
       • Time: 15 seconds
     • Annealing:
       • Temperature: 51°C
       • Time: 30 seconds
     • Extension:
       • Temperature: 72°C
       • Time: 90 seconds
   • Final Extension:
     • Temperature: 72°C
     • Time: 10 minutes
   • Hold:
     • Temperature: 4°C

4. Sample Loading:

   Load 25 μl of each PCR reaction mix, including the colony PCR reactions (for 10 colonies) and the positive control, into separate PCR tubes.

5. Thermocycler Run:

   Start the thermocycler program to run the colony PCR reactions.

6. Analysis:

   After the PCR run is complete, analyze the PCR products as needed (e.g., gel electrophoresis, DNA sequencing).

Objective:

To prepare a 1% agarose gel and run gel electrophoresis for DNA samples, including DNA templates and a ladder.

Materials:

• Agarose (0.3 g)
• 1X TAE buffer (30 ml)
• SYBR Safe dye (2 μl)
• Electrophoresis tank with combs
• DNA templates (Colony 1-10)
• 1 kb ladder
• Micropipettes and tips
• Electrophoretic power supply
• Power cord
• Labels

Procedure:

1. Preparation of 1% Agarose Gel

   1. Weigh 0.3 g of agarose and add it to 30 ml of 1X TAE buffer in a flask to make a 1% agarose gel.
   2. Heat the mixture until the agarose completely dissolves and the solution becomes clear.
   3. After dissolving, add 2 μl of SYBR Safe dye to the agarose solution and mix gently.

2. Electrophoresis Tank Setup

   1. Set the electrophoresis combs to be approximately 2 cm away from the cathode side of the gel.
   2. Ensure that the gel tank is clean and dry.

3. Pouring the Agarose Gel

   1. When the agarose solution reaches a temperature of approximately 60 degrees Celsius, pour it into the central part of the gel tank.
   2. Allow the gel to solidify by keeping it undisturbed at room temperature.

4. Adding 1X TAE Buffer:

   Pour 1X TAE buffer into the gel tank until the buffer level stands at 0.5-0.8 cm above the gel surface.

5. Well Formation:

   Gently lift the combs from the gel, ensuring that the wells remain intact for loading samples.

6. Sample Loading:

   Load the DNA samples into the wells as follows for both gels:

   • Well 1 (of both gels): 1 kb ladder (6 μl)
   • Wells 2-7 (gel 1): Colony 1-6 DNA templates (25 μl each)
   • Wells 2-5 (gel 2): Colony 7-10 DNA templates (25 μl each)

7. Electrophoresis

   1. Connect the power cord to the electrophoretic power supply.
   2. Set the electrophoresis parameters as needed (e.g., voltage, run time) and start the electrophoresis.
   3. Allow the electrophoresis to run until the DNA samples have migrated sufficiently to achieve the desired separation.

8. Visualization and Documentation

   1. After electrophoresis, remove the gel from the tank and place it on a UV transilluminator or gel documentation system.
   2. Visualize the separated DNA bands using UV light.

9. Data Recording:

   Document the gel image and record the positions and sizes of DNA bands.

10. Safety Precautions:

    Follow appropriate safety measures when working with UV light sources and SYBR Safe dye.

Objective:

To perform SDS-PAGE (Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis) for protein analysis.

Materials:

• Gel casting unit with glass plates and spacers
• Deionized water
• Ethanol
• 30% Acrylamide
• 1M Tris (pH 8.8)
• 10% SDS
• 10% APS(freshly prepared)
• TEMED
• 0.5M Tris (pH6.8)
• Ethanol (for gel surface overlay)
• Comb for creating wells in the gel
• Sample loading buffer
• Protein samples (Induced and Uninduced)
• Electrophoresis apparatus
• Coomassie Blue dye
• Destaining solution
• Plastic container
• Shaker

Procedure:

1. Preparation of Gel Casting Unit

   1. Clean the glass plates and spacers of the gel casting unit with deionized water and ethanol.
   2. Assemble the plates with the spacers on a stable, even surface.

2. Preparation of Resolving Gel

   1. Prepare the resolving gel solution using the following volumes for a 12.5% gel:

   2. Component	Resolving Gel
      MilliQ water	1.151mL
      30% Acrylamide	2.079mL
      1M Tris: pH 8.8	1.663mL
      10% SDS	50μl
      10% APS(freshly prepared)	50μl
      TEMED	5μl

   3. Pour the gel solution into the plates assembled with spacers. To maintain an even and horizontal resolving gel surface, overlay the surface with ethanol.
   4. Allow the gel to set for about 20-30 minutes at room temperature.

3. Preparation of Stacking Gel

   1. Prepare the stacking gel solution using the following volumes:
   2. Allow the gel to solidify by keeping it undisturbed at room temperature.

   3. Component	Stacking Gel
      MilliQ water	1.752mL
      30% Acrylamide	0.369mL
      0.5M Tris: pH 6.8	0.324mL
      10% SDS	25μl
      10% APS(freshly prepared)	25μl
      TEMED	3μl

   4. Discard the overlaid ethanol on the resolving gel.
   5. Add the 5% stacking gel solution until it overflows. Insert the comb immediately ensuring no air bubbles are trapped in the gel or near the wells.
   6. Allow the gel to set for about 20-30 minutes at room temperature.

4. Sample Preparation and Loading

   1. Add 1 mL of loading buffer to each of the samples (Induced and Uninduced) of 1 mL each and incubate at 99 degrees for 20 minutes.
   2. Load these samples onto the gel along with two ladder markers.
   3. Perform electrophoresis at 50V for 3 hours.

5. Staining and Destaining

   1. Carefully remove the gel and place it in a plastic container.
   2. Add Coomassie Blue dye to the gel and leave it overnight on a shaker.
   3. The next day, add a destaining solution and keep it for 4 hours.
   4. Observe the gel under white light.