Plasmid extraction:-

We used the Promega kit for plasmid extractions.

1. Add 600μl of bacterial culture to a 1.5ml microcentrifuge tube. (For higher yields and purity use the alternative protocol below to harvest and process up to 3ml of bacterial culture.)
2. Add 100μl of Cell Lysis Buffer (Blue), and mix by inverting the tube 6 times.
3. Add 350μl of cold (4–8°C) Neutralization Solution, and mix thoroughly by inverting.
4. Centrifuge at maximum speed in a microcentrifuge for 3 minutes.
5. Transfer the supernatant (~900μl) to a PureYield™ Minicolumn without disturbing the cell debris pellet.
6. Place the minicolumn into a Collection Tube, and centrifuge at maximum speed in a microcentrifuge for 15 seconds.
7. Discard the flowthrough, and place the minicolumn into the same Collection Tube.
8. Add 200μl of Endotoxin Removal Wash (ERB) to the minicolumn. Centrifuge at maximum speed in a microcentrifuge for 15 seconds.
9. Add 400μl of Column Wash Solution (CWC) to the minicolumn. Centrifuge at maximum speed in a microcentrifuge for 30 seconds.
10. Transfer the minicolumn to a clean 1.5ml microcentrifuge tube, then add 30μl of Elution Buffer or nuclease-free water directly to the minicolumn matrix. Let it stand for 1 minute at room temperature.
11. Centrifuge for 15 seconds to elute the plasmid DNA. Cap the microcentrifuge tube, and store eluted plasmid DNA at –20°C.

Restriction digestion protocol:-

We used the restriction digestion protocol from Promega corp.

1. In a sterile tube, assemble the following components in the order listed below. Mix gently by pipetting after every addition.

   Component	Volume (μl)
   Sterile, deionized water	16.3
   Restriction enzyme 10X buffer	2
   Acetylated BSA (10 μg/μl)	0.2
   DNA (1 μg/μl)	1.0
   Restriction enzyme (10 u/μl)	0.5
   Final volume	20

2. Mix gently by pipetting, close the tube and centrifuge for a few seconds in a microcentrifuge. Incubate at the enzyme’s optimum temperature for 1–4 hours.
3. Add loading buffer to a 1X final concentration and proceed to gel analysis.

Note: Overnight digestions are usually unnecessary and may result in DNA degradation. And for double digestion use the buffer that gives >=50% efficiency for both enzymes.

Agarose gel electrophoresis:-

Materials required:-Agarose solution ((0.5-7 kb bands )-1% of agarose [w/v]), 50X TAE stock, EtBr for DNA staining, 6X gel loading buffer, DNA ladder, sample

1. Mix agarose powder with 100 mL 1xTAE in a microwavable flask.
2. Microwave for 1-3 min until the agarose is completely dissolved (but do not overboil the solution, as some of the buffer will evaporate and thus alter the final percentage of agarose in the gel. Many people prefer to microwave in pulses, swirling the flask occasionally as the solution heats up.).
3. Let agarose solution cool down to about 50 °C (about when you can comfortably keep your hand on the flask), about 5 mins.
4. Add ethidium bromide (EtBr) to a final concentration of approximately 0.2-0.5 μg/mL.
5. Pour the agarose into a gel tray with the well comb in place
6. Place newly poured gel at 4 °C for 10-15 mins OR let sit at room temperature for 20-30 mins, until it has completely solidified.
7. Once solidified, place the agarose gel into the gel box (electrophoresis unit).
8. Fill gel box with 1xTAE (or TBE) until the gel is covered.
9. Carefully load a molecular weight ladder into the first lane of the gel. Then carefully load your samples into the additional wells of the gel.
10. Run the gel at 80-150 V until the dye line is approximately 75-80% of the way down the gel. A typical run time is about 1-1.5 hours, depending on the gel concentration and voltage.
11. Turn OFF power, disconnect the electrodes from the power source, and then carefully remove the gel from the gel box.
12. Using any device that has UV light, visualize your DNA fragments. The fragments of DNA are usually referred to as ‘bands’ due to their appearance on the gel.

DNA elution:-

We used the promega DNA concentration and elution kit for eluting out our DNA fragments. Note: Perform all centrifugation steps at 10,000 × g (14,000rpm).

1. Load and run your gel using an established protocol. DNA can be extracted from standard or low-melt agarose gels run with either TAE or TBE buffer.
2. Weigh a 1.5ml microcentrifuge tube for each DNA fragment to be isolated, and record the weight.
3. Visualize and photograph the DNA using a long-wavelength UV lamp and an intercalating dye such as ethidium bromide. To reduce nicking, irradiate the gel for the absolute minimum time possible. Excise the DNA fragment of interest in a minimal volume of agarose using a clean scalpel or razor blade. Transfer the gel slice to the weighed microcentrifuge tube, and record the weight. Subtract the weight of the empty tube from the total weight to obtain the weight of the gel slice.

   Note: The gel slice may be stored at 4°C or at –20°C for up to 1 week in a tightly closed tube under nuclease-free conditions before purification.

4. Add Membrane Binding Solution at a ratio of 10μl of solution per 10mg of agarose gel slice.
5. Vortex the mixture and incubate at 50–65°C for 10 minutes or until the gel slice is completely dissolved. Vortex the tube every few minutes to increase the rate of agarose gel melting. Centrifuge the tube briefly at room temperature to ensure the contents are at the bottom of the tube. Once the agarose gel is melted, the gel will not resolidify at room temperature.
6. Load sample onto a ReliaPrep™ Minicolumn seated in a Collection Tube and centrifuge for 1 minute.
7. Remove minicolumn and discard Collection Tube contents. Reseat the minicolumn into the same Collection Tube.
8. Note: For gel slices >350mg, continue to pass additional sample through the ReliaPrep™ Minicolumn until all of the sample has been loaded.
9. Add 200μl of Column Wash Solution (CWE) and centrifuge for 15 seconds. Remove minicolumn and discard the contents of the Collection Tube. Reseat the minicolumn into the same Collection Tube.
10. Wash with 300μl of Buffer B (BWB) and centrifuge for 15 seconds. Repeat wash with 300μl of Buffer B (BWB) and centrifuge again.
11. Remove minicolumn and discard the contents of the Collection Tube. Reseat the minicolumn into the same Collection Tube and centrifuge for 1 minute to dry the minicolumn. Then transfer minicolumn to an Elution Tube.
12. Pipet 15μl of Nuclease-Free Water or TE buffer (not provided) to the center of the minicolumn and centrifuge for 30 seconds.
13. For maximum recovery, repeat elution with an additional 15μl of Nuclease-Free Water or TE buffer.

Ligation:-

We used the ligation protocol from Promega corp. We recommend using a 1:1, or 1:3 molar ratio of vector:insert DNA when cloning a fragment into a plasmid vector.

PCR:-

We followed the protocols from Promega corp.

1. Thaw the PCR Master Mix at room temperature. Vortex the Master Mix and then spin it briefly in a microcentrifuge to collect the material in the bottom of the tube.
2. Add the following components in the given sequence:-

   Component	Volume	Final Conc.
   PCR Master Mix, 2X	25μl	1X
   upstream primer, 10μM	0.5–5.0μl	0.1–1.0μM
   downstream primer, 10μM	0.5–5.0μl	0.1–1.0μM
   DNA template	1–5μl	<250ng
   Nuclease-Free Water to final volume of	50μl	N.A.

3. Set the thermal cycler at:-
   1. Denaturation: 95oC, 2 minute (initial), subsequent 1 minute
   2. Annealing: 5oC below primer Tm, 1 minute
   3. Extension: For Taq DNA pol, 72-74oC, calculate time using 1kb per minute extension rate in cycles, final extension 5 minute

   Perform 25-30 cycles.

Overlapping PCR:-

1. PCR individual parts.
2. Prepare PCR mixture, without primers. Instead of a template, add your PCR parts. Use a large volume, i.e. 1/2 to 3/4 of the total PCR reaction. Make sure to use a molar ratio of ~1:1.
3. Add the two primers flanking the outer parts as you would in a normal PCR.
4. Rerun your PCR at 30 cycles, this time using an annealing temperature matching your flanking primers.

Note: Important: Gel-extract your overlap extension product, as this method can result in non-specific side products! PCR reaction will likely yield multiple bands (e.g. the fragments you started out with), as well as a smear around the desired band.

Ni-NTA chromatography:-

We provide this protocol from the Promega kit.

1. Pipet 700μl of bacterial culture or supernatant (if protein is secreted out) into a 1.5ml microcentrifuge tube. Add 70μl of the FastBreak™ Reagent/DNase I solution to the culture.
2. Resuspend the resin and allow it to settle. Once the resin has settled, use a wide-bore pipette tip to transfer 75μl of the HisLink™ Resin from the settled resin bed to the 1.5ml microcentrifuge tube. To successfully transfer resin, place the wide-bore pipette tip deep into the resin and pipet slowly to assure that a consistent amount of resin is drawn into the pipette. Allow the resin to resettle between each pipetting. Note: We recommend optimizing the amount of HisLink™ Resin used for low- (<1mg/ml) or high- (>1mg/sample) expressing proteins. For low-expressing proteins, less resin should be used; similarly, for high-expressing proteins, more resin per sample can be used.
3. Incubate the sample and resin for 30 minutes, mixing frequently on a rotating platform or shaker to optimize binding.
4. Place a Spin Column onto a Collection Tube (or a new 1.5ml microcentrifuge tube). Use a wide-bore pipette tip to transfer the lysate and resin from the original 1.5ml microcentrifuge tube in Step 3 to the spin column. If resin remains in the 1.5ml microcentrifuge tube, add HisLink™ Binding/Wash Buffer to the tube, then transfer the buffer and remaining resin to the spin column.
5. Centrifuge the spin column with the collection tube for 5 seconds or until the liquid clears the spin column
6. To save the flowthrough, remove the spin column from the collection tube and transfer the flowthrough from the collection tube to a new 1.5ml microcentrifuge tube. Otherwise, discard the flowthrough.
7. Place the spin column back onto the collection tube. Add 500μl of HisLink™ Binding/Wash Buffer to the spin column, then cap the spin column. Centrifuge for 5 seconds or until the Binding/Wash Buffer clears the spin column. Discard the flowthrough. Repeat for a total of two washes.
8. Take the spin column off the collection tube and wipe the base of the spin column with a clean absorbent paper towel to remove any excess HisLink™ Binding/Wash Buffer.
9. Place the spin column onto a new 1.5ml microcentrifuge tube. Add 200μl of HisLink™ Elution Buffer. Cap the spin column and tap or flick it several times to resuspend the resin. Wait 3 minutes. Note: HQ-tagged proteins may elute with a lower concentration of imidazole (50–100mM) compared to polyhistidine-tagged proteins.
10. Centrifuge the spin column and microcentrifuge tube at 14,000rpm for 1 minute to collect the eluted protein.

Bradford assay:-

Materials required:- Bradford dye concentrate, Phosphate-buffered saline (PBS), Protein standard (see Step 1), Protein to be assayed

1. Prepare a series of protein samples for a standard curve. Use PBS for making the dilutions. The standard curve will be linear between ~20 and 150 μg in 100 μl. Use a protein with similar properties as your sample (i.e., if testing antibody concentrations, use purified antibody for the standard). If the composition of your sample is unknown, use antibody.
2. Prepare serial dilutions of the test samples in PBS the same way as for the protein standard.
3. Dilute the Bradford dye concentrate using 1 part dye to 4 parts H2O. Filter if any precipitate develops.
4. Add 5 ml of diluted dye to each sample. Allow the color to develop for at least 5 minutes but for no longer than 30 minutes. The red dye will turn blue as it binds protein. Read the absorbance of each sample at 595 nm.
5. Centrifuge the spin column with the collection tube for 5 seconds or until the liquid clears the spin column
6. Generate a standard curve using the absorbance reading of the protein standard, and use it to calculate the concentrations of the unknown samples. BSA gives a value about twofold higher than its weight for Bradford dye-binding assays.

SDS-PAGE:-

Materials required:-

• Buffers for discontinuous buffer system
• 4X Stacking gel buffer (0.5 M Tris-HCl at pH 6.8)
• 4X Resolving gel buffer (1.5 M Tris-HCl at pH 8.8)
• 10X Running buffer
• 2X SDS-PAGE sample buffer
• n-Butanol (H2O-saturated)
• Methanol
• Ammonium persulfate solution (10%)
• TEMED
• Protein solution(s) or pellets of cellular proteins
• SDS (10%) stock solution
• 30%T (2.6%C) Acrylamide stock solution

Pouring a Slab Gel

1. Clean the glass plates.
   1. Soak the glass plates in 2% PCC-54 cleaning solution for 3-24 hours.
   2. Rinse the plates with tap water thoroughly and then once with distilled H2O.
   3. Dry the glass plates with clean tissue paper, and then clean them with Kimwipes soaked in methanol. Dry the plates in the air.
2. Assemble the gel-casting unit.
   1. Form the gel sandwich by assembling the spacers and two glass plates in the clamps.
   2. Align the bottom part of the spacers and two glass plates at the same level, and then tighten the clamp.
   3. Place the gel sandwich onto the casting stand.
   4. Insert the Teflon sample application comb, and mark the glass plate at a level ~1.0-1.5 cm below the bottom of the comb teeth.
3. Pour the resolving gel.
   1. Prepare the appropriate resolving gel mixture using the recipes in Table 1. Make sure that the solution is well mixed before adding the TEMED.
   2. Use a 10-ml pipette to transfer the mixture to the glass-plate sandwich up to the marked level (marked in Step 2iv).
   3. Carefully overlay the gel with an ~2-mm-deep layer of H2O or H2O-saturated n-butanol or isopropanol solution. This prevents air from reaching the gel, which inhibits polymerization of the acrylamide, and ensures that the gel surface is flat.
   4. After polymerization is complete (~30 min), pour off the overlaying H2O, and carefully remove any remaining liquid with filter paper without damaging the gel surface. If the gel is overlaid with n-butanol (or isopropanol), drain the overlay liquid, and then wash the gel surface with H2O. Polymerization of the gel is evidenced by a clear refractive index change that can be seen between the gel and the overlay liquid.
4. Pour the stacking gel.
   1. Select an acrylamide concentration for the stacking gel, and make the appropriate mixture, using the recipes in Table 2. Make sure that the solution is well mixed.
   2. Carefully overlay the resolving gel with the stacking gel solution until the height of the stacking gel is ~2.0-3.0 cm.
   3. Insert the Teflon comb into this solution, leaving 1.0-1.5 cm between the top of the resolving gel and the bottom of the comb. Make sure that no air bubbles are trapped beneath the teeth of the comb. Insert the comb into the stacking gel at an angle to reduce the chance of trapping air bubbles under the comb’s teeth. Trapped bubbles can be released by gently tapping on the glass plate near the trapped bubbles.
   4. Allow the stacking gel mixture to polymerize for ~2 hours. Refractive index changes around the comb indicate that the gel has set. It is useful at this stage to mark the positions of the bottoms of the sample wells on the glass plates with a marker pen.

Alternatively, remove trapped air bubbles by squirting running buffer across the bottom edge of the gel through a long bent needle or hooked Pasteur pipette. The gel is now ready to receive protein samples. Preparation of Samples Carry out either Step 7 or Step 8.

5. Carefully remove the sample comb from the stacking gel, and assemble the cassette in the electrophoresis apparatus according to the manufacturer’s instructions.
6. Fill the top reservoir with running buffer ensuring that the buffer fully fills the sample loading wells, and look for any leaks from the top tank. If there are no leaks, fill the bottom tank with running buffer, then tilt the apparatus to dispel any bubbles caught under the gel.
7. Prepare protein solutions:
   1. Mix the protein solution with 2X SDS-PAGE sample buffer in a 1:1 ratio. Although under ideal conditions the binding ratio for SDS and polypeptide is 1.4 g of SDS per gram of polypeptide, to ensure that enough SDS is present, the concentration of protein in the final solution should not be higher than 10 μg/μl.
   To load the entire protein sample onto the gel, bear in mind that the sample volume (i.e., protein solution and sample buffer) should not exceed the volume of the wells
   2. Heat the samples in a heat block or water bath for 2 minutes at 95°C to denature the proteins and ensure the maximum amount of SDS binding to the proteins. Allow the samples to cool to room temperature. Remove insoluble materials by centrifugation.
8. Prepare total cellular protein samples:
   1. Loosen the prepared cell pellet by vortexing the pellet briefly.
   2. Add 2X SDS-PAGE sample buffer directly to the cell pellet and vortex. The resultant cellular protein lysate is highly viscous. The amount of sample buffer to add varies depending on the cell line under study. However, a good amount to use is 100 μl of 2X sample buffer per 1 × 105 cells. The optimal amount must be determined empirically. To load the entire protein sample onto the gel, bear in mind that the sample volume (i.e., protein pellet and sample buffer) should not exceed the volume of the wells.
   3. Centrifuge the cellular protein lysate at 100,000g for 20 minutes, and collect the supernatant. Protein concentration for a certain number of cells differs slightly for different cell lines. If the protein concentration exceeds 10 μg/μl, add SDS powder. Addition of SDS to 5% final concentration will not interfere with the electrophoretic separation.
   4. Heat the samples in a heat block or water bath for 2 minutes at 95°C to denature the proteins and ensure the maximum amount of SDS binding to the proteins. Allow the samples to cool to room temperature. Remove insoluble materials by centrifugation.

Running a Discontinuous Slab Gel

9. Use a pipette and gel-loading pipette tips to load the samples into the sample well.
10. Connect the power supply to the electrophoresis apparatus with the anode (+) linked with the bottom reservoir and the cathode (-) connected to the upper reservoir.
11. Pass a constant voltage of 200 V at 8°C (with cooling system connected), or 90 V at room temperature, through the gel until the Bromophenol Blue dye front reaches the bottom of the gel. This will take ~6-8 hours at constant 200 V and ~16-18 hours at constant 90 V.
12. Turn off the power supply, and disconnect the electrodes. Remove the gel plates from the apparatus, and carefully remove a spacer. Use the spacer to gently pry the gel plates apart, leaving the gel stuck to one plate.
13. Visualize the proteins using an appropriately sensitive staining method like western blot.

Western blot:-

Note: The antibodies used should be according to the purpose of the project.

1. Run SDS-PAGE gel, and then Western transfer the protein samples to nitrocellulose (NC) membrane for immunoblot analysis.
2. After transfer, transfer the membrane to western-blot tray, briefly wash the NC membrane with distilled water.
3. (Optional) Visualize the proteins on the membrane by Ponceau’s staining.
4. Wash off the red stain with distilled water.
5. Block the membrane with 5-10ml blocking buffer (made by 5% non-fat milk in 1xPBST) for 30 minutes at R/T.
6. Dilute the primary antibody with blocking buffer according to the suggested dilution factor on datasheet.
7. Remove the blocking buffer and add enough diluted primary antibody to cover the membrane.
8. Incubate the membrane with primary antibody for 1hr at R/T. (Note: Or you can do overnight incubation at 4C, make sure you cover the western-blot tray to prevent excessive evaporation). To prevent uneven coverage, the western-blot tray can be rocked on a rocker platform.
9. Collect the primary antibody and store them at 4C for up to two weeks. (If you would like to store them longer, you can freeze the diluted antibody at –20C. Remember frequent freezing and thawing will gradually decrease the antibody titer.)
10. Briefly wash the membrane with 1xPBST once to remove any excessive primary antibody.
11. Add enough 1xPBST to cover the membrane and leave the Western-blotting tray on a rocker platform.
12. Wash the membrane for 15 minutes. (Note: If the background is high, repeat this step for two to three times.), turn on the developer during the wash time.
13. Dilute HRP-conjugated secondary antibody with blocking buffer (1:1000 or higher dilution is usually good for Goat anti-mouse-HRP).
14. Incubate the membrane with secondary antibody for 30 minutes to 1hr.
15. Wash the membrane with 1xPBST for 15 minutes, and then 3 times (5 min/time).
16. Prepare the chemiluminescence development substrate mixture by mixing equal amount of solution1 and 2 (Normally 1ml will be enough for one membrane).
17. Prepare a plastic saran film, lay the film on a flat surface, and dispense 1ml of substrate mixture for one membrane on the plastic saran film.
18. Use a forceps to take washed the blot from the western-blotting tray, flip it, lay on the substrate mixture, and then incubate for 1 to 5 minutes. (Note: To avoid air bubbles, always lay the blot by touching one edge first.)
19. Remove excess Chemiluminescence Reagent and wrap the membrane in plastic. Place inside X-ray cassette.
20. Expose to film and develop.

Cell line culture:-

1. Observe cells to see how confluent they are, whether the cells are alive, whether the cells are contaminated, and whether the cells have the correct morphology. After cell culture reaches 80-85% confluence, subculture is conducted.
2. Remove media from dish.
3. Wash with 10 ml of 1X PBS.
4. Add 5 ml of Trypsin and trypsinize for 3 min at 37C. Whack hard – you should see the cells coming down. (Important: never overtrypsinize the cells, so work quickly)
5. Add 5 ml of media and use it to rinse the dish to detach the cells off (45 times). (The serum in the media will neutralize the trypsin)
6. Spin down at 1000rpm for 35 min at room temperature. Aspirate supernatant.
7. Add 15 ml of media to 15ml tube containing cell pellet, and pipette up and down to mix.
8. Add 15 ml of media to each new 150mm x 25mm tissue culture dish (we split 1 dish into 34 dishes)
9. Add 5mls of cell culture containing media to each tissue culture dish. Make sure the media covers the entire area of the dish. Put the cells into 37C with 5% CO2.

MTS assay:-

We mention the protocol from the Promega kit.

1. Thaw the CellTiter 96® AQueous One Solution Reagent. It should take approximately 90 minutes at room temperature, or 10 minutes in a water bath at 37°C, to completely thaw the 20ml size
2. Pipet 20μl of CellTiter 96® AQueous One Solution Reagent into each well of the 96-well assay plate containing the samples in 100μl of culture medium.
3. Note: We recommend repeating pipettes, digital pipettes or multichannel pipettes for convenient delivery of uniform volumes of CellTiter 96® AQueous One Solution Reagent to the 96-well plate.
4. Incubate the plate at 37°C for 1–4 hours in a humidified, 5% CO2 atmosphere. Note: To measure the amount of soluble formazan produced by cellular reduction of MTS, proceed immediately to Step 4. Alternatively, to measure the absorbance later, add 25μl of 10% SDS to each well to stop the reaction. Store SDS-treated plates protected from light in a humidified chamber at room temperature for up to 18 hours. Proceed to Step 4.
5. Record the absorbance at 490nm using a 96-well plate reader.

Annexin-5, PI staining – Flow cytometry:-

Materials required:-

1. Cultured cells, Toxicant -TRAIL-Smac Fusion Peptide
2. Phosphate buffered saline (PBS; Fischer or Sigma)
3. Annexin binding buffer
4. Annexin V–conjugated FITC (annexin V–FITC), typically at 25 μg/ml 1 mg/ml propidium iodide (PI) in PBS
5. Orbital shaker
6. 50-μm diameter nylon filter sieves, 5-ml snap-cap polypropylene tubes
7. Flow cytometer with FL-1 (FITC) and Fl-2 (PI) channels

Prepare and treat cells:-

1. Treat cells with the desired toxicant (see Critical Parameters for suggested reference toxicants). Typically, use 1 million cells per treatment group (i.e., for each time or concentration point). Set up controls as follows:
   1. For the PI positive control, treat a separate aliquot or dish of cells with a toxicant at a concentration known to disrupt plasma membrane integrity.
   2. For the annexin V positive control, treat a separate aliquot or dish of cells with a toxicant known to induce apoptosis.
   3. For the no-stain negative control, do not expose the cells in this aliquot or dish to any kind of treatment.
2. Be sure to include a separate set of controls for each experiment.
3. After incubation for a predetermined time, remove the toxicant and wash cells with PBS as follows:
   1. If adherent cells are used, wash the monolayers twice with 1 ml of PBS.
   2. If cellular suspensions are used or if cells have been released from the monolayers, centrifuge 5 min at 4000 × g, remove supernatant, and resuspend pellet in 1 ml PBS.
4. Add 600 μl of annexin binding buffer.
5. Add 2-5 μl of 25 μg/ml annexin V–FITC to each sample and to the annexin V positive control. Do not add annexin V–FITC to the no-stain negative control or PI positive control. Add 25 μl of 1 mg/ml PI to each sample and to the PI positive control. Do not add PI to the no-stain negative control or annexin V positive control.
6. Incubate the cells for 10 min at room temperature on an orbital shaker at 50 rpm in the absence of light.
7. Wash the cells to remove unbound annexin V–FITC and PI as follows:
   1. If adherent cells are used, aspirate buffer and add 1 ml of binding buffer without annexin V–FITC or PI. Incubate the cells for 10 min on an orbital shaker at 50 rpm. Shield cells from light at all times (i.e., cover with foil). Decant solution and repeat the wash and 10-min incubation in binding buffer two times. Add 500 μl annexin-binding buffer and gently remove the cells by scraping with a rubber policeman. Disperse the cells by pipeting, and filter the sample through a 50-μm nylon filter sieve into a labeled 5-ml snap-cap polypropylene tube.This step removes clumps and prevents clogging of the cytometer.
   2. If cell suspensions are used, centrifuge cells 5 min at 4000 × g, remove supernatant, and suspend the pellet in 1 ml of binding buffer. Repeat this process two times. After the final wash, add 0.5 to 1 ml of binding buffer to obtain 1 × 106 cells/ml.

Measure Annexin V and PI staining:-

8. Analyze the cells using flow cytometry with the proper channels for FITC (typically FL-1) and PI (typically FL-2).
9. Read the no-stain negative control sample first. Adjust the forward and side scatter settings and the Fl-1 and Fl-2 compensations to produce plots
10. Read the annexin V positive control. *An increase in fluorescence intensity on the Fl-1 channel will be observed. The pattern should be similar to that displayed in R4 and LR on Figure 12.8.3C and D, respectively. *A separate population may not be seen if adherent cells are used. Rather, a definite shift will be observed (see R4 Figure 12.8.3C). *If no shift in fluorescence intensity is seen along the Fl-1 channel, then the amount or concentration of annexin V–FITC added may be too low. Alternatively, the agent used to induce apoptosis was inactive.
11. Read the PI positive control. *An increase in fluorescence intensity on the Fl-2 channel should be observed.. *The PI positive controls are typically similar for both cellular suspensions and adherent cells released using a rubber policeman. *If no shift in fluorescence intensity is seen along the Fl-2 channel, the amount or concentration of PI added may have been too low. Alternatively, the agent used to induce necrosis may be inactive
12. Adjust the Fl-1 (annexin V–FITC) and Fl-2 (PI) channel compensations. Determine the settings that result in the highest amount of signal for each marker with the least amount of fluorescent overlap (bleed-over). Read the controls and samples under these conditions. *If the compensation on the FL-2 (PI) channel is maximized, there is the possibility that the FL-1 (FITC) signal will be reduced resulting in a masking or loss of the annexin V–FITC signal. Similarly, if the compensation of the Fl-1 (FITC) channel is maximized, then the Fl-2 (PI) signal may be reduced. *Once the forward scatter, side scatter, and the Fl-1 and Fl-2 compensations are set for the no-stain negative control and the annexin V and PI positive controls, they should not be altered during the rest of the analysis. If a change is made in any of these parameters all controls must be re-verified and samples re-tested.

Analyze data:-

13. For all analyses, use the no-stain negative control to determine the absolute boundaries for both annexin V–FITC and PI positive cells (Fig 12.8.3A and B). Samples can be examined by quadrant, histogram, or region analysis. Quadrants work well for cell suspensions (Fig. 12.8.3B) while region analysis is better than quadrants for adherent cells removed from cultures by mechanical methods (Fig. 12.8.3A).
14. After determining the proper type of analysis (gates, histograms, or regions), perform statistics on every sample, including the no-stain negative control and annexin V and PI positive controls. Be sure to compare data generated with control cells to that of the no-stain negative control. If using region analysis for adherent cultures, subtract the no-stain negative control values from the actual samples. Failure to do so could result in artificially high values for cell death due to the basal events present in the R2, R3, and R4 regions of the no-stain negative control (Fig. 12.8.3A). These statistical analyses can be performed using the software supplied with the flow cytometer. Control cells will remain in the same region or quadrant as the no-stain negative control (denoted R1 and LL in Fig. 12.8.3A and B, respectively). Apoptotic cells are those with increased fluorescent intensity on the annexin-FITC axis (Fl-1 scale), denoted in R4 and LR on Fig. 12.8.3C and D, respectively. Necrotic cells are identified by their increased fluorescent intensity on the PI axis (Fl-2 scale), denoted in R2 and UL in Figure 12.8.3E and F, respectively. Cells may also be present in the R3 region or UR quadrant. This type of staining may indicate late apoptosis when membrane integrity is finally lost, or necrosis when annexin-V–FITC freely enters the cells and binds to intracellular phosphatidylserine. Ideally, the control levels of annexin V–FITC binding should be ≤10%, while PI staining should be less than 5%. These values can be cell-type dependent.

Annexin-5, PI staining – Flow cytometry:-

For TRAIL-binding assay, we first saturate cell receptors with anti-DR4/5 antibodies and then introduce TRAIL-Smac, and then subsequently we introduce HRP conjugated anti-His antibody (our construct has a His tag). In the next experiment, we introduce only the TRAIL-Smac fusion contruct and then the anti-His antibody to ascertain whether TRAIL is bound to the DR4/5 receptor on the cell surface. This way we can check whether the TRAIL-Smac fusion construct binds only to the DR4/5 receptors. This protocol is also used to check for Smac internalization. Here we check for the presence of Smac inside the cells. The protocol for immunostaining that will be used is given below (this can be used for both the above mentioned assays):- Materials required:- Chamber slides, cover slips, or 12-well plates, Phosphate-buffered saline (PBS), Fixation solution: 1% Paraformaldehyde, in PBS, Blocking buffer: 5% FBS in PBS

1. Transfer a single cover slip into a 12-well plate. Then add 1 mL of 70% Ethanol into a well for 20 minutes at room temperature.
2. Wash quickly three times with PBS.
3. (Optional; for loosely attached cells) add 1 mL of 0.1 mg/mL Poly-D-lysine solution into a well for 15 minutes at room temperature. Wash quickly three times with PBS and let dry before plating cells.
4. Grow cultured cells on cover slips or in wells overnight at 37°C. At the time of fixation, cells should be ~70-80% confluent in single layer.
5. Rinse cells briefly in PBS.
6. Fix cells by incubation with freshly made 1% Paraformaldehyde in PBS for 10 minutes at room temperature.
7. Rinse three times quickly in PBS.
8. (Optional; for intracellular staining) add permeabilization solution and incubate at room temperature for 10 minutes. Then wash quickly three times in PBS.
9. Block samples in 1 mL of blocking buffer at room temperature for 30 minutes
10. Dilute the primary antibody to the recommended concentration/dilution in blocking buffer.
11. For 8-well chamber slides, add 200 µL per well. For 12-well plates, add 500 µL per well. Incubate two to three hours at room temperature or overnight at 4°C. If using conjugated antibodies, perform this step in the dark.
12. For surface staining, rinse 3 times quickly in PBS. For intracellular staining, quickly wash once followed by incubation with wash buffer for 5-10 minutes. Then quickly wash an additional two times. Note: If using primary antibodies directly conjugated to fluorochromes, then skip to step 17.
13. Prepare fluorochrome-conjugated or enzyme linked secondary antibody in blocking buffer according to the manufacturer’s specification data sheet, and add 200 µl per well to the 8-well chamber slides. For 12-well plates, add 500 µL per well.
14. Incubate the samples for one hour, at room temperature, in the dark.
15. For surface staining, rinse three times quickly in PBS. For intracellular staining, quickly wash once followed by incubation with wash buffer for 5-10 minutes, then quickly wash an additional two times.
16. Optional: To stain F-actin, prepare a working solution of Flash Phalloidin™ by diluting it 1:20-1:100 in PBS. Add 200 µL per well for an 8-well plate or 500 µL per well for a 12-well plate. Stain for 20 minutes at room temperature in the dark.
17. Apply anti-fade mounting medium to the cover slip.
18. Seal slides with nail polish.

Checking cleavage of fusion protein assay:-

We are sourcing this protocol from R&D systems, which is a vendor of MMP-3/9 and uPA enzymes (which are also overexpressed in most cancer cells) and our TRAIL-Smac fusion construct carries the protease sites for cleavage of the fusion protein resulting in subsequent Smac internalization. Materials required for MMP-3/9 based assay:- Assay Buffer: 50 mM Tris, 10 mM CaCl2, 150 mM NaCl, 0.05% (w/v) Brij­35, pH 7.5 (TCNB) Recombinant Human MMP­3 (rhMMP­3) Chymotrypsin, 1 mg/mL stock in 1 mM HCl Phenylmethyl Sulfonyl Fluoride (PMSF), 0.2 M stock in 2­Propanol Substrate protein, 2 mM stock in DMSO F16 Black Maxisorp Plate Fluorescent Plate Reader

1. Activate rhMMP­3 at 20 μg/mL in Assay Buffer containing 5 μg/mL Chymotrypsin.
2. Incubate reaction at 37 °C for 30 minutes.
3. Stop activation with 2 mM PMSF. Pre­warm the PMSF to 37 °C prior to adding to sample.
4. Dilute activated rhMMP­3 to 2.5 ng/μL in Assay Buffer.
5. Dilute Substrate to 20 μM in Assay Buffer.
6. In a plate load 50 μL of 2.5 ng/μL rhMMP­3, and start the reaction by adding 50 μL of 20 μM Substrate to wells. Include a Substrate
7. Blank containing 50 μL Assay Buffer and 50 μL of 20 μM Substrate.
8. Then perform SDS-PAGE for SmacN7 or TRAIL. The band for SmacN7 or TRAIL should shift from high molecular weight (fusion construct) to low molecular weight (cleaved protein).

Materials required for uPA based assay:- Assay Buffer: 50 mM Tris, 0.01% (v/v) Tween® 20, pH 8.5 Recombinant Human u‑Plasminogen Activator (uPA)/Urokinase (rhuPA) Substrate protein, 10 mM stock in DMSO F16 Black Maxisorp Plate Fluorescent Plate Reader

1. Dilute rhuPA to 1 ng/μL in Assay Buffer.
2. Dilute Substrate to 200 μM in Assay Buffer.
3. Load 50 μL of the 1 ng/μL rhuPA into a black well plate, and start the reaction by adding 50 μL of 200 μM Substrate. Include a Substrate Blank containing 50 μL Assay Buffer and 50 μL of 200 μM Substrate without any rhuPA.
4. Then perform SDS-PAGE for SmacN7 or TRAIL. The band for SmacN7 or TRAIL should shift from high molecular weight (fusion construct) to low molecular weight (cleaved protein).

Caspase-8 activation assay:-

This protocol has been sourced from Promega corp.

Note: Cells should be grown in multiwell plates that are adequate for cell culture and compatible with the luminometer being used.

1. Remove 96-well plates containing cells from the incubator, and allow plates to equilibrate to room temperature. Do not touch pipet tips to the wells containing samples if reusing tips to avoid cross-contamination.
2. Add 100μl of Caspase-Glo® 8 Reagent to each well of a white-walled 96-well plate containing 100μl of blank, negative control cells or treated cells in culture medium. Because of the sensitivity of this assay, be careful not to touch pipet tips to the wells containing samples to avoid cross-contamination. Cover the plate with a plate sealer or lid. Note: Lids must be removed before reading plate. Temperature fluctuations will affect luminescent readings.
3. Gently mix contents of wells using a plate shaker at 300–500rpm for 0.5–2 minutes. Incubate at room temperature for 30 minutes to 3 hours, depending upon the cell culture system.
4. Measure the luminescence of each sample in a plate-reading luminometer as directed by the luminometer manufacturer.

Caspase-3/7 activation assay:-

This protocol is from the Promega kit.

Note: Cells should be grown in multiwell plates that are adequate for cell culture and compatible with the luminometer being used. Directions are given for performing the assay in a total volume of 200μl using 96-well plates.

1. Remove 96-well plates containing treated cells from the incubator and allow plates to equilibrate to room temperature.
2. Add 100μl of Caspase-Glo® 3/7 Reagent to each well of a white-walled 96-well plate containing 100μl of blank, negative control cells or treated cells in culture medium. Because of the sensitivity of this assay, be careful not to touch pipet tips to the wells containing samples to avoid cross-contamination. Cover the plate with a plate sealer or lid. Note: If you are reusing pipet tips, do not touch pipet tips to the wells containing samples to avoid cross-contamination.
3. Gently mix contents of wells using a plate shaker at 300–500rpm for 30 seconds. Incubate at room temperature for 30 minutes to 3 hours, depending upon the cell culture system. The optimal incubation period should be determined empirically.

   Note: Temperature fluctuations will affect the luminescence reading. If the room temperature fluctuates, use a constant-temperature incubator.

4. Measure the luminescence of each sample in a plate-reading luminometer as directed by the luminometer manufacturer.

TUNEL assay:-

Materials required:- Cells of interest Citrate buffer (0.1% Triton X-100 in 0.1% sodium citrate) Cytotoxic agent of choice (In our project, it is the TRAIL-Smac fusion peptide)

Hoechst 33342 (Sigma-Aldrich)

In Situ Cell Death Detection Kit (Roche Diagnostics 11684795910)

Paraformaldehyde (PFA) (4%) in PBS Phosphate-buffered saline (PBS) ProLong Gold Antifade (Life Technologies)

1. Treat cells of interest with a cytotoxic agent (see Protocol: Triggering Apoptosis in Hematopoietic Cells with Cytotoxic Drugs (Crowley et al. 2015c) or Protocol: Triggering Death of Adherent Cells with Ultraviolet Radiation ([Crowley et al. 2015d]) and fix the cells in 4% PFA for 20 min at room temperature following incubation with the cytotoxic agent for the required period of time.

Adherent cells can be fixed in PFA directly. Nonadherent cells will need to be harvested and attached to the coverslip (see Protocol: Morphological Analysis of Cell Death by Cytospinning Followed by Rapid Staining [Crowley et al. 2015b]).

2. Wash the cells twice with PBS for 5 min each at room temperature. Pipette or aspirate off the PBS.

To avoid damaging the cells, do not touch them with the pipette

3. Permeabilize the cells by incubating them in citrate buffer for 2 min on ice.
4. Wash the cells twice with PBS for 5 min each at room temperature. Pipette or aspirate off the PBS.

To avoid damaging the cells, do not touch them with the pipette.

5. Prepare the terminal dUTP nick-end labeling (TUNEL) reagent by mixing 5 µL of enzyme solution with 45 µL of label solution per slide. Add 50 µL of TUNEL reaction mixture per slide and cover with Parafilm or a coverslip to ensure even coating of the cells. Incubate in the dark in a humidified incubator for 1 h at 37°C.

The TUNEL reagent must be freshly prepared before use. The reagent is light-sensitive and should be protected from light at all stages.

A negative staining control should be performed on a control slide using 50 µL of label solution without enzyme solution.

6. Remove the TUNEL reagent by aspiration. Wash the cells twice with PBS for 5 min each at room temperature.
7. Incubate the cells with Hoechst 33342 (100 ng/mL in PBS) in the dark for 15 min at room temperature.

Staining intensity can be changed by incubating the slides in stain for different periods of time.

8. Wash the cells twice with PBS for 5 min each at room temperature. Pipette or aspirate off the PBS.

To avoid damaging the cells, do not touch them with the pipette.

9. Rinse the cells with water.
10. Mount the cells with ProLong Gold Antifade reagent and cover with a coverslip.
11. Leave to dry overnight at room temperature. View the slides immediately using a fluorescence microscope or store in the dark.

TUNEL labeling indicates that DNA strand breaks have occurred in those cells. In contrast, cells stained only with Hoechst 33342 (and not TUNEL) do not have DNA strand breaks. The number of cells stained with both TUNEL and Hoechst 33342 (apoptotic) versus the number of cells stained with Hoechst 33342 alone (healthy) can therefore be used to determine the level of apoptosis in a sample. Images of cells can be captured using a fluorescence microscope with an attached camera and counted for graphical representation.

TUNEL-stained cells may also be assayed by flow cytometry. It is advisable to seek advice from expert users if analysis by flow cytometry is desired.

Growing B. longum in anaerobic conditions:-

This requires a special instrument called anaerobic chamber to generate the hypoxic environment needed to grow anaerobic bacteria.

1. A loop of inoculum will be transferred to 50 ml MRS media anaerobically, using a Anaerobic Chamber System and incubated for 24 hours at 37 degree C
2. This culture will be mixed with glycerol to 50 % (v/v) final Conc and stored at -80 degree C freezer
3. Inoculate 2ml from this glycerol stock into fresh MRS medium and incubate anaerobically at 37 degree C for 20 hours
4. Check OD of this culture and use it when the OD is between 0.4 to 0.6

Electrotransforming B. longum:-

1. Cultivate cells by using an overnight culture to inoculate fresh medium. Grow cells overnight at 37 °C. Dilute this culture 1:25 in fresh medium and cultivate at 37 °C until an O.D.695 of 0.2. Chill bacteria on ice.
2. Harvest by centrifugation.
3. Wash twice with 0.5 M sucrose
4. Resuspend in about 1/250 of the original culture volume of 1 mM ice-cold sucrose-citrate buffer, dispense in tubes and incubate for 3.5 hours at 4 °C.

Electroporation of cells:

1. Add 0.5-1.5 µg plasmid DNA to 80 µl of electrocompetent cells. Homogenize by gently mixing with pipette several times. Transfer mixture into prechilled cuvette.
2. Wipe moisture from the cuvette and insert the cuvette into the device.
3. Electroporation:

   mode	Prokaryote
   Voltage V	1,200 V
   Time	5 ms

4. Dilute with 800 µl outgrowth medium and incubate for 2.5 h at 37 °C.
5. Plate onto selective MRS agar plates; incubate anaerobically for 2-3 days at 37 °C.

Expected results: Transformation efficiency up to 9.4 x 104 transformants/µg of DNA.

References

Handling of rats for tumor model study:-

Induction of tumor – subcutaneous deposition in immunocompromised mice:-