Team:ASU/Experiments

Experiments

Agarose Gel Electrophoresis Protocol

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After extracting genomic DNA, running a gel will help determine sample quality. Electrophoresis is useful as a quantifying tool for the estimation of molecular weight and sample quality. ~30 uL aliquots of gDNA should be kept aside after the extraction to be used for this protocol. For Novogene, a g-DNA of ≥ 20 ng/uL is recommended with no degradation and no contamination.

Materials

Reagents and Buffers:

- DNA Samples

- 1X TAE buffer solution

- Agarose

- Gel loading dye

- DNA ladder

- DNA stain

Equipment:

- 500 mL Erlenmyer flask

- Microwave

- Graduated cylinder

- Spatula

- Gel casting tray

- Gel well comb

- Power supply and cables

- Gel box

- UV Light source

- Pipette tips

Procedure

1X TAE Solution Preparation:

1. Add 20 mL of TAE buffer to 980mL of ddH2O

Agarose gel concentration:

2. Choose appropriate gel concentration:

	Percent agarose gel (w/v)	DNA size resolution	Agarose to weigh out for 25 ml gel (g)
1	0.5	1,000 - 25,000	0.125
2	0.75	800 - 12,000	0.188
3	1.0	500 - 10,000	0.25
4	1.2	400 - 7,500	0.3
5	1.5	200 - 3,000	0.375
6	2.0	50 - 1,500	0.5

Agarose Gel Preparation:

3. Measure 0.25 g of agarose pwder (range of fragements 720 to 10,633 bps)

4. Mix agarose powder with 25 ml 1x TAE in a microwavable flask

5. Microwave for 1-3 min in small increments until agarose is completely dissolved (clear)

Make sure it doesn't boil over

6. Cool agarose to 65°C with careful swirling to promote even dissipation of heat

Measure using infrared thermometer

7. Prep gel-casting tray while the gel is cooling, make sure it is level

Select well comb size based on sample number and volume of sample you want ot run in the gel. Make sure the gel casting tray is tightly bound before pouring agarose. Make sure the tray is level so gel is evenly distributed.

8. Before casting the gel, add 5 μl SYBR Safe concentrate to the molten agarose and swirl to mix well

9. Pour the agarose into a gel tray with the well comb in place.

Pour slowly to avoid bubbles that disrupt gel.

Bubbles can be pushed to edges or popped with a pipette tip.

10. Let gel sit for 10-20 min at Room Temperature until completely solidified

11. Fill the gel box with enough 1X TAE buffer to fully submerge gel and wells need to be filled with buffer

12. Once gel is solidified, place casting tray and agarose gel into the gel box

13. After submerging the gel carefully remove the comb to expose the sample wells

The comb should be removed after submerging the gel in the TAE buffer, this prevents bubbles from being trapped in the sample wells

Loading DNA Samples and Running Gel

14. Add loading buffer to samples

How much loading buffer to add to samples:

	A	Sample
1	Sample (uL)	20
2	6X Loading buffer (uL)	4
3	Total to load	24

15. Use table below to specify sample organization

	A	B	C	D	E	F	G	H
1

16. Carefully load 20 uL of DNA ladder into the lanes as indicated in the table

Can be pipetted directly out of the stock ladder tube and does not need loading buffer

Pipette it slowly into the well without pushing too hard on the well or puncturing it. Carefully release the DNA ladder from the pipette tip right above the well.

17. Carefully, load 24 uL of DNA samples into corresponding wells

18. Connect power source to gel box and turn it on

Note: Black is negative, red is positive; DNA is negatively charged and runs toward the positive electrode

19. Watch the gel carefully in the first couple of minutes to ensure that the dyes are migrating in the correct direction

20. After electrophoresis is complete turn off power and disconnect electrodes from the power source

21. Using any UV light device, visualize your DNA fragments

Gel Visualization and Analysis:

22. Using any device that has UV light, visualize the DNA bands.

23. Carefully place the gel under UV light

24. Turn on the UV transilluminator

25. Use long wavelength UV and expose for little time to minimize DNA damage

26. Take a picture of data

27. Turn off UV transilluminator

28. Take out agarose gel and discard

Troubleshooting

29. Compare visualized data from gel electrophoresis to the gels below

30. In the figure above: (A) Trans2K™ Plus DNA Marker, (B) qualified sample, (C) degraded sample, (D) sample contaminated with RNA, (E) sample contaminated with protein. Red boxes denote areas of contamination.

31. Samples above show DNA degradation which would impact quality of prepared library

32. Samples above show RNA contamination. RNase should be added to DNA before shipping for exome sequencing.

33. Samples above show protein contamination. Protein purification through affinity chromatogrophy should be done before shipping DNA for exome sequencing

Safety Protocols

These protocols must be followed for the entire duration of the lab work:

1. All individuals in the laboratory must be wearing PPE at all times.

2. All individuals in the laboratory must have the appropriate training certifications required for the lab procedures, including hazardous waste, autoclave training, and general lab safety.

3. Store all gels and DNA in the proper plates at the correct temperature.

4. After the procedure, all organic waste should be disposed of in an organic waste container.

5. All remaining DNA and glassware should be autoclaved following the procedure, and then stored in the proper place.

6. Clean the workstation after the procedure is complete.

Ampicilin stock solution (100mg/ml)

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Store the ampicillin in aliquots at -20°C for 1 yr (or at 4°C for 3 mo)

Materials

- Ampicilin (sodium salt, m.w. 371.40)

Procedure

Make stock solution:

1. Dissolve 1 g of sodium ampicilin in sufficient ddH20 to make a final volume of 10 mL.

2. Prewash a syringe filter with 50ml of ddH2O.

3. Pass dissolved ampicilin through syringe filter.

4. Store in 500 ul aliquots in sterile 1.5 ml tubes labeled with Amp and date.

5. Store the ampicillin in aliquots at -20°C for 1 yr (or at 4°C for 3 mo).

Arsenic Solution

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Prepare 1000x (1g/L or 1000ppm) solution of Sodium arsenate dibasic heptahydrate for dilution.

Materials

- Sodium arsenate dibasic heptahydrate (CAS 10048-95-0)

- ddH2O

- Weigh boats

- Chemical fumehood

Procedure

Preparation of 1000x (1g/L or 1000ppm) solution

1. 1 g/L requires 1 g of arsenic. Divide the molar mass of total product (sodium arsenate) by the molar mass of arsenic.

312.01 (g/mol)/74.92(g/mol) = 4.16g

2. Add 4.16g of sodium arsenate to approximately 900ml of water.

3. Dissolve thoroughly and fill the remaining liquid (1000ml) with ddH2O.

Can cut volumes in half or even 1/10 to adjust total solution.

Bead Transformation Protocol

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Translated from Wannathong et al. 2016 "New tools for chloroplast genetic engineering allow the synthesis of human growth hormone in the green alga C. Reinhardtii. This protocol is for the transformation of recombinant DNA into the chloroplast of C. reinhardtii using glass beads.

Materials

- C. reinhardtii strain (CC-5168)

- Glass beads of 400-625 μm diameter

- TAP medium

- 5 ml test tube

- Vortexer (Vortex Genie II)

- TBP plates with appropriate selective antibiotic

- HSM Plates

- Top Agar (5%)

Procedure

Ahead of Time:

1. Aliquot 300mg of 400-625um diameter beads into 5-10ml test tubes. Cap and tape with autoclave tape.

Autocalve to sterilize.

2. Make top agar: 5% agar molten at 42 C.

Keep warm and in liquid state.

3. Prepare TBP and HSM plates.

4. Inoculate and culture 200-400mL of CC-5168 in TAP media until log phase has been reached (approximately 2*10^6 cells).

Day 1:

5. Aliquot out around 6mL of top agar into individual 15mL falcon tubes.

Ensure they remain liquid and do NOT solidify. Only remove from bath for brief periods of time.

Pellet cells by centrifugation and resuspend in 6ml of media.

7. Transfer 300ul of cells to sterile test tube containing 300mg of glass beads.

8. Add 5-10ug of miniprepped DNA.

We added 20uL of DNA.

9. Vortex on maximum speed for 15 seconds.

10. Pipette 300uL of vortexed cell/DNA mixture from test tube into 6mL of top agar (at 42C).

11. Invert gently to mix.

12. Plate top agar on HSM or TBP plate.

For each plasmid, one plate of each was made. Same top agar was used. All 6mL are used for one plate.

13. At room temperature (25C) Grow plates at dim light (~2 μE/m2/s) overnight (approximately 15 hrs).

14. Transfer to moderate light (~50 μE/m2/s) the next day. Grow for up to 3 weeks.

Be patient and check in regularly, they can take over a month to grow.

15. Restreak on selective media to confirm transformation.

Safety Protocols:

These protocols must be followed for the entire duration of the lab work.

1. All individuals in the laboratory must be wearing PPE at all times.

2. All individuals in the laboratory must have the appropriate training certifications required for the lab procedures, including hazardous waste, autoclave training, as well as general lab safety.

3. Store all algal cells and DNA in the proper plates at the correct temperature.

4. Ensure that the gene gun is properly set up and sterilized before and after use.

5. After the procedure, all organic waste should be disposed of in an organic waste container.

6. All remaining DNA and glassware should be autoclaved following the procedure, and then stored in the proper place.

7. Clean the workstation after the procedure is complete.

Chlamydomonas Arsenic Uptake

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This protocol fully outlines the experiments relating to the uptake of arsenic by Chlamydomonas.

Materials

Reagents:

- TAP media

- 10% nitric acid

Equipment:

- ICP-MS

- Shaker table

- Beakers with baffles for C. reinhardtii growth

- Light source

- Hemocytometer

- Falcon tubes

- Centrifuge

- Filters

Procedure

Inoculate a Culture of Chlamy:

1. Fill a beaker with 25 mL of TAP

2. Use a sterile toothpick to smear a few colonies from a plate.

3. Wet the side of the beaker by gently swirling the TAP, and spin the stick against the wet side of the beaker to transfer the colonies into the media and break up any clumps.

4. Gently swirl the tube and allow to shake at 200 rpm at room temperature with light until it reaches confluency. Continue diluting with additional TAP media until you reach a confluent volume of over 40 mL. This tends to take 2-3 days.

Pellet and Resuspension of Chlamy in Arsenate Solution (Day 1):

5. Count the cells in the culture using a hemocytometer.

6. Calculate the volume of the culture required to achieve a cell count of 5*10^5 cells.

7. Transfer this volume of cells into a falcon tube six times. Label three of them 50 ppb and the other three as 500 ppb, along with the name of your construct.

8. Spin down the cells in a centrifuge at 21C at 7100 rpm for 5 minutes.

9. Decant the supernatant, retaining the pellet.

10. Resuspend the pelletted cells in 1 mL of TP media.

11. Add the following volumes of resuspended cells, TP media, bicarbonate, and arsenic solution in order to obtain the same concentration (106 cells/mL) in each sample:

	Tube Label	Volume of Cells	Volume of TP	Volume of Bicarbonate	Volume of Arsenic Solution (1g/L)	Total Volume
1	50 ppb	1 mL	38.2 mL	0.8 mL	2 uL	40 mL
2	500 ppb	1 mL	38.2 mL	0.8 mL	2 uL	40 mL

12. Immediately remove 10 mL of solution from each of the three beakers of biological triplicates (3 samples for 500 ppb and 3 samples for 50 ppb).

13. Count the cells in each sample using a hemocytometer and record them:

	Tube Label	Concentration from hemocytometer
1	50 ppb (B1)
2	50 ppb (B2)
3	50 ppb (B3)
4	500 ppb (B1)
5	500 ppb (B2)
6	500 ppb (B3)

14. Spin these down in falcon tubes at 7,100 rpm for 5 minutes at room temperature.

DNA Gel Extraction and Nanodrop

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After running a gel electrophoresis to ensure your DNA is the size you wanted, you can excise it from the gel to use in later experiments. This protocol extracts the DNA and quanitifies it using a Nanodrop.

Materials

Reagents:

- Monarch® DNA Gel Extraction Kit (NEB #T1020)

- >95% ethanol

- Agarose gel with DNA to be extracted

- Nuclease free water

Materials:

- Scale

- Incubator

- Centrifuge (16,000 x g)

- Pipettes and tips

- Microcentrifuge tubes

- Scapel

- Nanodrop

- Kimwipes

Procedure

Buffer Preparation (only performed once):

1. Add ethanol to Monarch DNA Wash Buffer prior to use.

2. For one 50-prep kit add 20 ml of ethanol to 5 ml of Monarch DNA Wash Buffer.

DNA Extraction:

3. Excise the DNA fragment to be purified from the agarose gel using a razor blade, scalpel or other clean cutting tool. Use care to trim excess agarose.

4. Transfer the excision to a 1.5 ml microcentrifuge tube and weigh the gel slice.

5. Add 4 volumes of Monarch Gel Dissolving Buffer to the tube with the gel slice (e.g., 400 μl buffer per 100 mg agarose). If the gel slice is >150 mg, consider reducing the amount of Gel Dissolving Buffer to 3 or 3.5 volumes to minimize the guanidine salt present in the workflow.

Note: If the volume of the dissolved sample exceeds 800 μl, the loading of the sample onto the column should be performed in multiple rounds to not exceed the volume constraints of the spin column.

6. Incubate the sample between 37–55°C (typically 50°C), inverting periodically until the gel slice is completely dissolved (generally 5–10 minutes).

Note: For DNA fragments > 8 kb, an additional 1.5 volumes of water should be added after the slice is dissolved to mitigate the tighter binding of larger pieces of DNA (e.g., 100 mg gel slice: 400 μl Gel Dissolving Buffer: 150 μl water). Failure to dissolve all the agarose will decrease the recovery yield due to incomplete extraction of the DNA and potential clogging of the column by particles of agarose.

7. Insert the column into collection tube.

8. Load sample onto the column.

9. Spin for 1 minute, then discard flow-through.

10. Re-insert column into collection tuvbe.

11. Add 200 μl DNA Wash Buffer and spin for 1 minute. Discarding flow-through is optional.

12. Repeat wash (Steps 10 and 11).

13. Transfer column to a clean 1.5 ml microfuge tube. Use care to ensure that the tip of the column has not come into contact with the flow-through. If in doubt, re-spin for 1 minute before placing into a clean microfuge tube.

14. Add ≥ 6 μl of DNA Elution Buffer to the center of the matrix. Wait for 1 minute, and spin for 1 minute to elute DNA.

Note: Typical elution volumes are 6–20 μl. Nuclease-free water (pH 7–8.5) can also be used to elute the DNA. Yield may slightly increase if a larger volume of DNA Elution Buffer is used, but the DNA will be less concentrated. For larger size DNA (≥ 10 kb), heating the elution buffer to 50°C prior to use can improve yield. Care should be used to ensure the elution buffer is delivered onto the matrix and not the wall of the column to maximize elution efficiency.

NanoDrop Measurement:

15. Prepare the Nanodrop by turning it on (power button is in the back) and selecting 'dsDNA' from the measurement panel.

16. Apply 5 µL of nuclease free water to the bottom pedestal.

17. Lower the upper pedestal arm to form a liquid column; let it sit for approximately 2-3 minutes.

18. Gently wipe away the water from both the upper and lower pedestals with a clean kimwipe.

19. Add 1 µL RNAse/DNAse free water (use the same water that you eluted with). This will serve as the "blank".

20. Lower the bar and let the machine read.

21. Lift arm when told.

22. Wipe both pedestals gently with a clean kimwipe.

23. To read a sample, load the sample by adding 1 uL to the pedestal. Allow it to form a "bubble" on the pedestal.

24. Lower arm and allow the machine to perform a read.

25. Lift arm when told.

26. Record data in table below:

	Sample name	ng/ul	260/280	260/230	Notes
1
2
3
4
5

27. Wipe with a kimwipe.

28. Repeat steps 23-27 for as many samples as you have.

29. When your experiment is finished, select 'End Experiment'. You may save your data on a flashdrive by following the instructions on the Nanodrop, or you may record them on the table below.

30. Clean pedestal and close arm before leaving.

31. Record your activity in the logbook.

Chemical Transformation

∨

Allows chemically competent cells to take in plasmid DNA. This is a useful way to amplify the concentration of your recombinant DNA for extraction later.

Materials

NEB 5-alpha Competent E. Coli cells

SOC

LB+ampR plates

Procedure

Transformation

1. Thaw a tube of NEB 5-alpha Competent E. coli cells on ice for 10 minutes.

2. Add 2 µl containing 1 pg-100 ng of plasmid DNA to the cell mixture (25ml). Carefully flick the tube 4-5 times to mix cells and DNA. Do not vortex.

3. Place the mixture on ice for 30 minutes. Do not mix.

4. Heat shock at exactly 42°C for exactly 30 seconds. Do not mix.

5. Place on ice for 5 minutes. Do not mix.

6. Pipette 950 µl of room temperature SOC into the mixture.

7. Place at 37°C for 60 minutes. Shake vigorously (250 rpm) or rotate.

8. Warm selection plates to 37°C.

9. Mix the cells thoroughly by flicking the tube and inverting, then perform several 10-fold serial dilutions in SOC.

10. Spread 50-100 µl of each dilution onto a selection plate and incubate overnight at 37°C. Alternatively, incubate at 30°C for 24-36 hours or 25°C for 48 hours.

Gene Gun Transformation

∨

Protocol acquired from Dr. Redding's lab. This protocol outlines the biolistic shooting of DNA into the microalgae Chlamydomonas reinhardtii (Chlamy). Using Tungsten particles (W-part) to shoot the DNA directly into the chloroplast of the Chlamy allows for more accurate take-up of the plasmid than electroporation.

Materials

- 0.05 g Tungsten Particle (W-Part)

- Sterile Eppendorf tube

- 0.05 mL 50% glycerol

- Tris-acetate-phosphate

- 500mL bottle(s) or 50mL conical tube(s)

- Plates for Chlamy

- 2 μl of 0.5 μg/μl (500 ng/μL) DNA

- 25 μl 100 mg/ml W-part

- 25 μl 2M CaCl2 (filter sterilized)

- 10 μl 0.1M spermidine

- Sterile Filter Holders

- Hemocytometer

- Water Bat Sonocator

- Cetrifuge, 10 min at 3.5k

Gene Gun: Helium gas with pressure between 400-500 psi, vacuum pump, lab timer, stolenoid valve

Procedure

Ahead of Time:

1. Sterilize filter holders in 70% ethanol (v/v in Millipore water).

2. Let sit for at least an hour and then let dry in running clean bench.

3. Make Tungsten particle (W-part) suspension: in glove box, tap out about 50mg (.05g) W-part into sterile pre-weighed Eppendorf tube.

4. Remove from the glove box and weigh the tube with the W-part in it.

5. Add sterile 50% glycerol so that W-parts are 100 mg/mL and then pipette up and down.

6. Sonicate thoroughly (about 20 sec in water bath sonicator at full power).

7. Make sure you have sterilized whatever you are going to spin the cells down in.

To sterilize centrifuge bottles do not put them in the autoclave with their lids on because as they cool, a vacuum is drawn which seals the lid on and warps the bottles. Instead, wrap the lids in foil, cover the tops of the bottles in foil and autoclave them separately

8. Ensure you have enough TBP plates: Shoot at least 4 plates for each construct.

Algal Cells:

9. Grow cells in TAP (Tris-acetate-phosphate) to 1-2 x 10^6 cells/ml.

10. Keeping everything sterile, centrifuge for 10 min at 3.5 K in 500 ml bottles or in 50 ml conical tubes at room temperature.

11. Re-suspend in a small volume of TAP (or other) media and look at the cells in the hemocytometer.

12. Count the cells and plate approximately 10^7 cells per plate. Let dry at least 5 min.

Preparing DNA:

13. Mix in order in a microfuge tube:

i.) 2 μl 0.5 μg/μl (500 ng/μL) DNA

ii.) 25 μl 100 mg/ml W-part

iii.) 25 μl 2M CaCl2 (filter sterilized)

iv.) 10 μl 0.1M spermidine (filter sterilized→the aliquots in the freezer are already filter-sterilized; base, not HCL!)

14. Let sit on ≥ 5 min (up to an hour).

15. Spin briefly (2 sec.) in microfuge

16. Remove 25 μl supernatant.

17. Vortex

18. Sonicate until solution is uniformly colored - no clumps

19. Leave on ice

Set-Up System:

20. Check to make sure there is He gas pressure in the cylinder

21. Set up vacuum pump and turn on.

22. Set up system inside hood:

plug in lab timer

plug in solenoid valve into timer.

set timer to 0.1 seconds.

23. Connect hose to regulator and solenoid valve

24. Open He gas cylinder and set regulator to desired pressure (between 400-500 psi).

25. Try some test shots to make sure system is working

(blast sterilized filter holderst to remove residual ethanol if necessary).

26. Wipe down inside of chamber with 75% ethanol to sterilize.

Shooting:

27. Re-vortex DNA-W

28. Briefly sonicate DNA-W solution

Solution should appear evenly colored.

29. Quickly after sonocation, apply 8μl DNA-Q solution on a sterile filter holder.

30. Screw filter holder onto underside of chamber top.

31. Place pre-labeled (strain and plasmid) Petri plate on lab jack at the proper height (11 cm from tip to agar).

32. Close chamber (make sure it is centered on gasket).

33. Turn on vacuum; make sure the release valve is closed and has a sterile filter on it.

34. When vacuum is at 25, hit the timer trigger to shoot.

35. Open release valve and turn off pump.

36.Open chamber and put top on plate and then para-film plate.

37. Suggestion from Marburg: Store in the dark for 24hours before transferring to the light.

38. Store plates in the light. Check every few days (up to a month) for transformed colonies.

Safety Protocols:

These protocols must be followed for the entire duration of the lab work.

1. All individuals in the laboratory must be wearing PPE at all times.

2. All individuals in the laboratory must have the appropriate training certifications required for the lab procedures, including hazardous waste, autoclave training, as well as general lab safety.

3. Store all algal cells and DNA in the proper plates at the correct temperature.

4. Ensure that the gene gun is properly set up and sterilized before and after use.

5. After the procedure, all organic waste should be disposed of in an organic waste container.

6. All remaining DNA and glassware should be autoclaved following the procedure, and then stored in the proper place.

7. Clean the workstation after the procedure is complete.

8. All individuals operating/setting up the helium tank must be certified in ASU’s EHS pressurized gas safety training.

Polymerase Chain Reaction (PCR)

∨

This protocol delineates the design of the polymerase reaction and thermal cycles.

Materials

- Forward/Upstream Primer

- Reverse/Downstream Primer

- Template DNA

- Polymerase Master Mix

- Nuclease Free Water

Procedure

PCR Reaction Mix:

1. Determine size of region of interest

-Identify gene of interest and record sequence in Benchling including any modifications (i.e., terminator sequence)

-Determine primer size and location including any modifications (i.e., homology regions or restriction sites)

-Add the gene of interest fragment size and primer fragment sizes together to estimate total length (or just use benchling tool)

2. Pick a polymerase; Taq polymerase for non-cloning amplification, or high fidelity polymerase for cloning

-Different polymerases have different temperature ranges and times for the reaction.

3. Set up reaction mixture in PCR tube with template DNA, upstream and downstream primers, polymerase master mix and nuclease free water

PCR Reaction Mix

	A	B	C	D
1	Component	25 µl reaction	50 µl reaction	Final Concentration
2	10 µM Forward Primer	0.5 µl	1 µl	0.2 µM (0.05–1 µM)
3	10 µM Reverse Primer	0.5 µl	1 µl	0.2 µM (0.05–1 µM)
4	Template DNA	variable	variable	less than 1,000 ng
5	Taq 2X Master Mix	12.5 µl	25 µl	1X
6	Nuclease-free water	to 25 µl	to 50 µl

4. Calculate annealing temperature and extension times and determine thermal cycles

-Determine GC content and primer concentration to calculate annealing temperature (https://tmcalculator.neb.com/#!/main)

-For extension time, add approximately 30-60 seconds of time per kilobase in amplified region of interest

-For all other times, use higher recommended time

-Ex. Taq 2x thermal cycler conditions

	A	B	C	D
1	STEP	TEMP	TIME	TIME
2	Initial Denaturation	95°C	3 minutes
3	30 Cycles	95°C (Denature)		15-30 seconds, 15-60 seconds, 1 minute per kb
4		45-68C (annealing)		15-60 seconds
5		68C (Extension)		1 minute per kb
6	Final Extension	68°C		5 minutes
7	Hold	4-10°C		Can stay as long as necessary. Store in 4C fridge or -20C for long-term

5. Store samples in 4C fridge or freeze in -20C fridge for long-term.

6. Dilute sample with loading del and run on .7% - 1% agarose gel to confirm amplification

Gibson Assembly

∨

Gibson Assembly Protocol from Unvieristy of Toronoto iGEM 2018 & from New England BioLabs Inc. (NEB): Gibson Assembly® Protocol (E5510). Gibson Assembly is the basic method of cloning that introduces an insert into a plasmid without the use of Type IIS restriction enzymes.

Materials.

- Milli-Q Water (deionized H2O)

- DNA fragments (gBlocks/inserts): Store at -20°C in TE for up to 24 months or nuclease-free H2O for up to 1 month

- Gibson Assembly Master Mix (2x): Store at -20°C

- Vectors (plasmids)

- Ice

Equipment

- Thermocycler

- Ice Bucket

- -20°C Fridge

- Autocalve Safe Disposable Bags

- Autoclave

Additional Notes:

- Efficiency of assembly decreases as the number or length of inserts increases.

- The mass of each insert can be measured using the NanoDrop instrument, with absorbance at 260 nm or estimated from agarose gel electrophoresis followed by ethidium bromide staining.

- Yields will be best when the different inserts are present in equimolar concentrations.

- Inserts to be assmbled should not have stable single-stranded DNA secondary structure, such as a hairpin or a stem-loop, or repeated sequences, as this would directly compete with the required single-stranded annealing/priming of neighboring assembly fragements.

- The Gibson Cloning Master Mix consists of three different enzymes within a single buffer. Each enzyme has a specific and unique function for the reaction:

- T5 Exonuclease: creates single-strand DNA 3’ overhangs by chewing back from the DNA 5’ end. Complementary DNA fragments can subsequently anneal to each other.

- Phusion DNA Polymerase: incorporates nucleotides to “fill in” the gaps in the annealed DNA fragments.

- Taq DNA Ligase: covalently joins the annealed complementary DNA fragments, removing any nicks and creating a contiguous DNA fragment.

Procedure

Calculations:

1. Calculating Optimal pmols of Each Insert:

Equation: Equation: pmols = (1000×weight in ng) / (length in bp×650 Da)

50 ng of 5000 bp dsDNA is about 0.015 pmols

Online Tools: NEBioCalculator: https://nebiocalculator.neb.com/#!/ligation

NEB Reccomendation:

0.02–0.5 pmols when 1-2 inserts are being assembled into a vector (or 2-3 fragments are being assembled)

0.2–1.0 pmoles when 3–5 inserts are being assembled into a vector (or 4-6 fragments are being assemble

2. Reaction Component Table

	A	B	C	D
1	Assembly Type	2-3 Fragment Assembly (vector + 1-2 inserts)	4-6 Fragment Assmebly (vector + 3-5 inserts)	Positive Control**
2	Total Amount of Fragements	0.02–0.5 pmols* (x μL)	0.2–1 pmols* (x μL)	10 μL
3	2X Gibson Assembly Master Mix	10 μL	10 μL	10 μL
4	Milli-Q water	(10-x) μL	(10-x) μL	0
5	Total Volume	20 μL***	20 μL***	20 μL

3. *Optimized cloning efficiency is 50–100 ng of vectors with 2–3 fold of excess inserts. Use 5 times more of inserts if size is less than 200 bps. Total volume of unpurified PCR fragments in Gibson Assembly reaction should not exceed 20%.

4. **Makes enough positive control reagents for five experiments.

5. ***If greater numbers of fragments are assembled, additional Gibson Assembly Master Mix may be required.

Procedure

6. Set up the reaction on ice using the reaction component table above.

7. Incubate samples at 50°C in a thermocycler for 15 minutes for 2-3 fragment assembly or 60 minutes for 4-6 fragment assembly.

Note: Reaction times less than 15 minutes are generally not recommended. Extended incubation times (up to 4 hours) have been shown to improve assembly efficiencies in some cases. Do not incubate the reaction overnight.

8. Following incubation, store samples on ice or at –20°C for subsequent transformation.

9. Transform competent ​E. coli​ cells with 2μL of the assembly reaction, following the transformation protocol.

Safety and Waste Disposal

10. Cleanup any tubes or micropipette tips by disposing in autoclave safe disposable bags and autoclave according to autoclave brand guidelines but cycle time must be set for a minimum of 30 minutes @ 1210C, 15 psi

Glycerol Stock Solution Protocol

∨

Bacterial glycerol stocks are important for long-term storage of plasmids.

Materials

- Srcew Cap Tubes

- 50% glycerol

- Overnight culture

- 70% Ethanol

- Autoclave-safe disposable bags

Procedure

Preparing Glycerol Stock Solution:

1. Label screw cap tubes.

2. Add 500 μL of the overnight culture to 500 μL of 50% glycerol in a 2 mL screw top tube or cryovial.

Note: Make the 50% glycerol solution by diluting 100% glycerol in dH20.

Note: Snap top tubes are not recommended as they can open unexpectedly at -80°C.

3. Gently mix.

4. Freeze the glycerol stock tube at -80°C.

5. Rinse glassware with 70% ethanol, dispose of in culture waste container, and clean with soap and water.

6. Dispose of any used screw cap tubes by rinsing contents with 70% ethanol.

7. Cleanup any tubes or micropipette tips by disposing in autoclave safe disposable bags and autoclave according to autoclave brand guidelines but cycle time must be set for a minimum of 30 minutes @ 121C, 15 psi.

Golden Gate Assembly

∨

This protocol is intended for use with the Type IIS restriction enzyme SapI. Golden Gate Assembly is a method of molecular cloning that can easily combine multiple fragments of DNA inserts. Using Type IIS restriction enzymes, the resulting product is "scarless" and does not retain the recognition sites, making it essentially irreversible. This protocol was taken from the DTU Denmark 2019 iGEM team.

Materials

- T4 DNA Ligase

- 10X T4 DNA ligase buffer

- Sapl Enzyme

- Receiver Plasmid

- DNA fragment with Sapl Overhangs

- ddH2O

- PCR Tubes

Procedure

Reaction Set-Up:

1. Add the following reagents to a PCR tube:

- 0.5 uL of T4 DNA ligase

- 2 uL of 10X T4 DNA ligase buffer

- 0.5 uL of SapI enzyme

- 100 ng of receiver plasmid

- equimolar amounts of inserts

- ddH2O for a total volume of 20 uL

2. Mix the contents of the PCR tubes gnetly by flicking.

3. Place the tube in a thermocycler.

Thermocycler Program:

	A	B	C
1	Step	Temperature	Time
2	1: Activation of Sapl	37°C	5 minutes
3	2: Activation of T4 Ligase	16°C	5 minutes
4	Repeat steps 1 and 2 for 25 cycle
5	3: Inactivation of Sapl	65°C	20 minutes
6	4: Inactivation of T4 ligase	85°C	10 minutes
7	5: Hold	4°C	Hold

4. Cleanup any tubes or micropipette tips by disposing in autoclave safe disposable bags and autoclave according to autoclave brand guidelines but cycle time must be set for a minimum of 30 minutes @ 121C, 15 psi.

Midi Prep

∨

Protocol taken from Dr. Kevin Redding's copy of the QIAGEN Plasmid Purification Handbook. The purpose of a midi prep is to extract large quantities of your desired plasmid. Source: https://www.protocols.io/view/culturing-chlamydomonas-reinhardtii-bkrskv6e?step=2

Materials

• QUIAGEN Midiprep kit
• Buffer P1
• Buffer P2
• Buffer P3
• QIAGEN tip 100
• QIAfilter cartridge
• LB+AmpR medium
• E. Coli + Plasmid
• Ethanol
• Sorvali SS-34 Centrifuge
• Tubes for Centrifuge
• Incubator
• Pipettes + tips
• Falcon Tubes
• TE Buffer

Procedure

MIDI prep with TIP 100

1. Pick a single colony from a sleective plate and innoculate a starter culture of 2-5 mlLB medium with selective antibiotic (amp).

2. Incubate for ~8 hrs at 37C with vigorous shaking; Volume of flask should be at least 4 times the volume of the culture

3. Dilute the starter cultuer 1/500 to 1/1000 into selective LB medium.

4. Inoculate 25mL of medium.

5. Grow at 37C for 12-16 hours with vigorous shaking. Volume of flask should be at least 4 times the volume of the culture. Culture should look cloudy but not too dense.

6. Harvest the bacterial cells by centrifugation at 6000x g for 15 min at 4C; 6000x g corresponds to 6000 rpm in Sorvall GSA or GS3.

7. Carefully decant all supernatant; If needed, freeze cell pellet at -20 C (or -80??)

8. Resuspend the bacterial pellet in 4mL of Buffer P1. Ensure pellet is completely resuspended by pipetting up and down or mixing gently.

9. Add 4mL of Buffer P2 to the lysate.

10. Mix immediately but genetly by inverting the tube 4-6 times. Proceed directly to the next step. Do not incubate on ice.

11. Pour the lysate into the barrel of the QIAfilter cartridge. Incubate at room temperature for 10 minutes. Do not insert the plunger.

12. Equilibrate a QIAgen tip 100 by applying 4 mL buffer QBT and allow the column to empty by gravity flow.

13. Remove the cap from the QIAfilter cartridge outlet nozzle.

14. Gently insert the plunge into the QIAfilter midi cartridge and filter the cell lysate into the previously equilibrated QIAgen tip.

15. Allow the cleared lysate to enter the resin by gravity flow.

16. Wash the QIAgen tip with 10 mL buffer QC two times.

17. Elute the DNA with 5 mL of buffer QF into a new tube.

18. Precipitate the DNA by adding 3.5 mL of room temperature isopropanol to the eluted DNA.

19. Mix and centrifuge immediately at greater than 15,000 g for 30 minutes at 4C.

20. Carefully decant the supernatant. Do not decant the pellet.

21. Air dry the pellet for 5-10 minutes.

22. Redissolve the DNA in 200 uL of 1X TE solution.

LB Agar Stock

∨

LB agar is used to make LB plates to grow bacterial cultures on. This should be kept very sterile and will solidify quickly as it is an agar solution.

Materials

- Bacto-tryptone (VWR 90000-286)

- Yeast extract (VWR IC10330380)

- NaCl (VWR BDH9286-2.5KG)

- Agar (VWR 97064-336)

- 1L sterile bottle with screw cap

- NaOH solution (1M)

- Sterile plates

Procedure

LB Agar Stock Recipe:

1. Weigh out 10 grams of bacto-tryptone. Add it to a sterile bottle.

2. Weigh out 5 grams of yeast extract and add it to the same bottle.

3. Weigh out 10 grams of NaCl and add it to the same bottle.

4. Weigh out 12 grams of agar and add it to the same bottle.

5. Add ddH2O up to one liter.

6. Mix well by shaking the bottle.

7. Once the reagents have fully dissolved, adjust the pH to 7.0 by adding small amounts of sodium hydroxide solution (NaOH).

8. If you are adding antibiotic to the media, follow the "Ampicillin stock solution" protocol to make solution (if you are using Ampicillin) and pour in 500 µL of antibiotic stock solution into your cooled (55°C) LB agar, stirring well, before pouring your plates.

9. If using shortly, place in a water bath set for 55°C. If storing for longer, keep at room temperature and autoclave before use.

Pouring Plates:

10. Carefully pour a thin layer of your LB agar solution into the bottom of the plates (the bottom will be smaller). Work quickly but carefully to avoid making bubbles or letting the agar solidify. Feel free to wear heat resistant gloves.

11. Allow the plates to cool with the lids propped to prevent condensation falling back onto the agar.

12. Once the agar is solid (about 20 minutes), stack the plates with the bottom side facing up and store them in the fridge. Ideally, you should stack the plates and put them back in their plastic packaging for storage.

Miniprep and Nanodrop Protocol

∨

Miniprep kit is used to elute DNA from a sample. The handbook dictates that this protocol is designed for the purification of up to 20 μg of high-copy plasmid DNA from 1–5 ml overnight cultures of E. coli in LB medium, but we are using 3mL specifically.

Materials

- Miniprep kit (QIAprep Spin Miniprep Kit (250) PN: 27106)

- LB amp bacterial plate

- LB amp liquid media

- 37°C incubator

- 37°C shaker incubator

- Microcentrifuge

- Vortex mixer

- 100% Ethanol

- Microcentrifuge tubes 2mL sterile

- Microcentrifuge tubes 1.5mL sterile (DNase free)

- Pipettes (1000 μL and 200 μL) and tips

Procedure

Steak out bacterial plate (Day 1):

1. Streak out bacteria on LB amp plate using the pattern below with three sterile pipette tips:

Start overnight (O/N) culture (Day 2):

2. Label culture tube with plasmid names.

3. Add 6mL of LB amp media.

4. Pick a single colony with a pipette tip from a freshly streaked selective plate and drop the pipette tip into correspondingly labeled tube.

5. Incubate for 12–16h/overnight at 37°C on shaking incubator.

Collect cell pellet (Day 3):

6. Pellet 1.5 ml bacterial culture (not to exceed 15 OD units) by centrifugation at 16,000 x g (~13,000 RPM) for 30 seconds. Discard supernatant.

Qiagen Miniprep Kit

7. Resuspend pellet in 200 μl Plasmid Resuspension Buffer (B1) (pink). Vortex or pipet to ensure cells are completely resuspended. There should be no visible clumps.

8. Lyse cells by adding 200 μl Plasmid Lysis Buffer (B2) (blue/green). Invert tube immediately and gently 5–6 times until color changes to dark pink and the solution is clear and viscous. Do not vortex! Incubate for one minute or less.

9. Neutralize the lysate by adding 400 μl of Plasmid Neutralization Buffer (B3) (yellow). Gently invert tube until color is uniformly yellow and a precipitate forms. Do not vortex! Incubate for 2 minutes.

10. Clarify the lysate by spinning for 5 minutes at 16,000 x g.

11. Carefully transfer supernatant to the spin column and centrifuge for 1 minute. Discard flow-through.

12. Re-insert column in the collection tube and add 200 μl of Plasmid Wash Buffer 1. Plasmid Wash Buffer 1 removes RNA, protein and endotoxin. (Add a 5 minute incubation step before centrifugation if the DNA will be used in transfection.) Centrifuge for 1 minute.

13. Add 400 μl of Plasmid Wash Buffer 2 and centrifuge for 1 minute.

14. Transfer column to a clean 1.5 ml microfuge tube. Use care to ensure that the tip of the column has not come into contact with the flow-through. If there is any doubt, re-spin the column for 1 minute before inserting it into the clean microfuge tube.

15. Add ≥ 30 μl DNA Elution Buffer to the center of the matrix. Wait for 1 minute, then spin for 1 minute to elute DNA.

Measuring with Nanodrop:

16. Prepare the Nanodrop by turning it on (power button is in the back) and selecting 'dsDNA' from the measurement panel.

17. Apply 5 µL of nuclease free water to the bottom pedestal.

18. Lower the upper pedestal arm to form a liquid column; let it sit for approximately 2-3 minutes.

19. Gently wipe away the water from both the upper and lower pedestals with a clean kimwipe.

20. Add 1 µL RNAse/DNAse free water (use the same water that you eluted with). This will serve as the "blank".

21. Lower the bar and let the machine read.

22. Lift arm when told.

23. Wipe both pedestals gently with a clean kimwipe.

24. To read a sample, load the sample by adding 1 uL to the pedestal. Allow it to form a "bubble" on the pedestal.

25. Lower arm and allow the machine to perform a read.

26. Lift arm when told.

27. Record data in table below:

	Sample name	ug/ul	260/280	260/230	Notes
1
2
3
4
5

LACK OF DATA IN THIS TABLE (CONSULT)

28. Wipe with a kimwipe.

29. Repeat steps 35-39 for as many samples as you have.

30. When your experiment is finished, select 'End Experiment'. You may save your data on a flashdrive by following the instructions on the Nanodrop, or you may record them on the table below.

31. Clean pedestal and close arm before leaving.

32. Record your activity in the logbook.

Safety Protocols:

Reminder: These are to be completed both before and after the experiment has been completed. This is to ensure safety for both for the people doing the protocl as well as the people in the laboratory.

1. Any person in the laboratory must have proper PPE. This includes appropriate clothes, lab coats, and goggles. They must also be trained on the proper certifications/training that is needed for the appropriate lab and the equipment they will be using.

2. Store purified DNA cultures in cold refrigerator.

3. Dispose of pippetes, kim wipes in appropriate places.

4. Place leftover media, buffers etc in storage for continued use.

5. Autoclave proper matererial including dishware, tubes etc.

6. Clean up environment.

Promega DNA Purification for Chlamy

∨

This protocol was originally taken from Promega's "Wizard® Genomic DNA Purification Kit Technical Manual" and was adapted by Dr. Kevin Redding's lab to be specific to Chlamydomonas. This protocol is being used to extract genomic DNA from Chlamydomonas reinhardtii, specifically the CC-4388 strain. The extracted DNA will be verified with primers in PCR.

Materials

- **Some Materials From Promega Wizard Genomic DNA Purification kit**

- 150 uL Nuclei Lysis solution

- 0.75 uL RNase A solution

- 50 uL Protein Precipitation Solution

- 15 uL room temperature 70% Ethanol

- 25 uL DNA Rehydration Solution

- Microcentrifuge tube(s) (For each extract 2)

- -80 freezer

- Vortex

- Centrifuge

- Wood stick(s)

- Overnight 4 C storage

Procedure

Extract Chlamydomonas DNA (nucleic and Chloroplast)

1. Add 150ul of Nuclei Lysis solution to a microcentrifuge tube.

2. Add a small blob of fresh Chlamydomonas reinhardtii cells from a plate using a wooden stick. The blob should not be larger than a small unpopped popcorn kernel. Too many cells or cells that are too old (more than 10 days) will affect yield.

3. Place the tubes in the -80C freezer for 60 minutes.

4. Incubate cells at 65C for 15 mins.

5. Add 0.75ul of RNas A solution.

6. Incubate at 37C for 15 mins.

7. Cool sample to room temp for 5 mins. ** begin perparing the 150 ul isopropanol into microcentrifuge tubes now for step 5**

8. Add 50ul of Protein Precipitation Solution.

9. Vortex IMMEDIATELY after adding Protein Precipitation Solution for 30 seconds.

10. Centrifuge at 13000-16000xg for 3 mins.

11. Transfer supertantant to a clean microcentrifuge tube containing 150 ul of room temperature isopropanol.

12. Mix thoroughly by inversion.

13. Centrifuge 13000-1600xg for 1 min.

14. Aspirate supernatant and add 150ul of room temperature 70% ethanol.

Caution: Pellet will be loose!! Do not lose it! If the pellet is too loose, repeat centrifugation before aspirating

15. Centrifuge 13000-16000xg for 1 min

16. Aspirate the ethanol. Remove as much as possible CAREFULLY.

17. Air-Dry pellet for 15 mins.

18. Add 25ul of DNA Rehydration Solution.

19. Rehydrate at 65C for one hour or overnight at 4C.

Restriction Digestion Verification of Plasmid (pASapl)

∨

Restriction enzyme digestion is commonly used in molecular cloning techniques and used to quickly check the identity of a plasmid by a diagnostic digest. Restriction enzyme digestion uses naturally occurring enzymes that cleave DNA at specific sites or sequences to prepare DNA for analysis. The results of the restriction digest can be evaluated by gel electrophoresis in which DNA is separated by molecule length. Using CutSmart buffer as it is optimal for BsaI-HF. THe pASapI cloning vector - 6,622 bp plasmid, with BsaI sites at 1,326 bp, 3,930 bp, and 6,512 bp. See table of expected fragments from single and double digests in Notes section.

Materials

- Plasmid DNA (pASapl)

- Appropriate restriction enzyme:

Bsal-HF (NEB R3733) comes with buffer: rCutSmart

- DDh2O

- Pipette tips

Procedure

Bsal restriction digest:

	Reagent	Stock Concentration	Amount Required	Volume
1	pDNA	146.1 ng/ul	1 ug	6.84 ul
2	Bsal	20,000 U/ml	10 U	1 ul
3	rCutSmart	10x	1x	3 ul
4	ddH2O	-	Up to 30 ul	19.16 ul

1. Label a 1.5 mL tube.

2. Add water.

3. Add buffer.

4. Add plasmid DNA.

5. Add restriction enzyme.

6. Mix gently by flicking tube.

7. Incubate tube at 37 ˚C for 1 hour.

Notes:

8. To visualize the results of your digest, conduct gel electrophoresis:

	Fragment	No digest	Bsal digest
1	1	6,622	2,604
2	2	None	2,582
3	3	None	1,436

Safety Protocols:

Reminder: These are to be completed both before and after the experiment has been completed. This is to ensure safety for both for the people doing the protocol as well as the people in the laboratory.

1. Any person in the laboratory must have proper PPE. This includes appropriate clothes, lab coats, and goggles. They must also be trained on the proper certifications/training that is needed for the appropriate lab and the equipment they will be using. In this case, they will need hazardous waste, autoclave training, as well as general lab safety.

2. Store DNA in cold refrigerator within the proper agar plates.

3. Dispose of any used pippete tips in the proper waste container (not regular trash).

4. Organic waste product that does not need to be autoclaved such as reagents that can be discarded should be placed in the organic waste container.

5. DNA, glassware, waste, that needs to be autoclaved should be bound up neatly and placed into the autoclave. After it has been finihsed place the following material in the apporiate places.

6. Clean up of environment.

Resuspending gBlocks Gene Fragments

∨

Once received from IDT, you must resuspend, quantify, and calculate copy number of your gBlocks before using them in assembly.

Materials

Reagents:

- Microcentrifuge tubes

- RNAse/DNAse free water

- Kimwipes

Equipment:

- Microcentrifuge

- Vortex

- Incubator

- Nanodrop

Procedure

Resuspension:

1. Before opening the tube, spin it down in a microcentrifuge for 3–5 seconds to ensure the DNA is in the bottom of the tube.

2. Add molecular grade water to reach a final concentration of 10 ng/µL.

3. Vortex briefly.

4. Incubate at approximately 50°C for 15–20 min.

5. Briefly vortex and centrifuge.

6. Verify the final concentration (see below).

Quantification using Nanodrop:

7. Prepare the Nanodrop by turning it on (power button is in the back) and selecting 'dsDNA' from the measurement panel

8. Apply 5 µL of nuclease free water to the bottom pedestal.

9. Lower the upper pedestal arm to form a liquid column; let it sit for approximately 2-3 minutes.

10. Gently wipe away the water from both the upper and lower pedestals with a clean kimwipe.

11. Add 1 µL RNAse/DNAse free water (use the same water that you resuspended with). This will serve as the "blank".

12. Lower the bar and let the machine read.

13. Lift arm when told.

14. Wipe both pedestals gently with a clean kimwipe.

15. To read a sample, load the sample by adding 1 uL to the pedestal. Allow it to form a "bubble" on the pedestal.

16. Lower arm and allow the machine to perform a read.

17. Lift arm when told.

18. Record data in table below:

	Sample name	ug/ul	260/280	260/230	Notes
1
2
3
4
5

EMPTY TABLE (CONSULT)

19. Wipe with a kimwipe.

20. Repeat steps 15-19 for as many samples as you have.

21. When your experiment is finished, select 'End Experiment'. You may save your data on a flashdrive by following the instructions on the Nanodrop, or you may record them on the table below.

22. Clean pedestal and close arm before leaving.

23. Record your activity in the logbook.

Calculating copy number:

24. If it is necessary to dilute the resuspended gBlocks Gene Fragment to a specific copy number/µL, use the molecular weight and fmol/ng conversions for each gBlocks fragment which are provided on the spec sheet provided with the fragment.

25. Calculate the copy number/µL with the following equation:

(C) (M) (1 x 10–15 mol/fmol) (Avogadro’s number) = copy number/µL

Where C is the current concentration of the gBlocks Gene Fragment in ng/µL, and M is the molecular weight in fmol/ng, as provided on the spec sheet.

Safety Protocols:

Reminder: These are to be completed both before and after the experiment has been completed. This is to ensure safety for both for the people doing the protocl as well as the people in the laboratory.

1. Any person in the laboratory must have proper PPE. This includes appropriate clothes, lab coats, and goggles. They must also be trained on the proper certifications/training that is needed for the appropriate lab and the equipment they will be using.

2. Discard any disposable material used ie Kim Wipes in the appropriate bins.

3. Store DNA gblocks in refrigerator for future use.

4. Autoclave all tubes, equipment used in experiment.

5. Remember to be careful using electronics within the lab. They must be separated from equipment/lab material and careful of any hazards.

Safety With GMOs

∨

Careful handling with genetically engineered microorganisms and proper disposal will reduce health and safety risks while maintaining a minimally contaminated environment.

Materials

Proper Personal Protective Equipment (PPEs):

- Gloves

- Goggles/Eyewear

- Labcoat

- Closed-Toed Shoes

- Long Pants

Cleaning Materials and Disposal:

- 70% Ethanol

- Kimwipes

- 10% Bleach

- Dish Soap

- Water

- Autoclave Safe Disposable Bags

- Autoclave

Procedure

Maintaining a Safe Environment:

1. Wear PPEs at all times when handling or around microorganisms or hazardous/toxic materials

2. Disinfect bench surfaces, lab areas and equipment with ethanol before and after use.

3. Dispose of all plastic waste including tubes, pipettes and micropipette tips in appropriate autoclave safe disposable bags.

4. Cleanup any tubes or micropipette tips by disposing in autoclave safe disposable bags and autoclave according to autoclave brand guidelines but cycle time must be set for a minimum of 30 minutes @ 1210C, 15 psi.

5. Dispose of volumes of microbial waste or culture by mixing with 10% bleach and store in safe plastic or glass waste container with a mark for disposal.

6. Rinse any glassware containing antibiotics or culture/biological waste with 70% ethanol and store in safe plastic or glass waste container with a mark for disposal. Clean the glassware with soap and water and autoclave to remove any microbial contaminants.

7. Dispose of any broken glassware in appropriate contianers, and clean any biological waste with 70% ethanol.

Western Blot

∨

Part of the process of purifying recombinant proteins, Western Blot checks the tagged constructs to verify expression of our target genes.

Materials

• 50 mL falcon tubes
• Wash buffer (1-10 mL):
• 50mM Tris-HCl pH 7.5
• 10mM EDTA
• 10x Protease Inhibitors (PI):
• 100mM EDTA
• 10mM benzzmidine - HCl
• 500 ug/mL pepstatin A
• 200 ug/mL leupeptin
• 100 uM E64
• 100 mM e-amino caproic acid, freeze
• XCell SureLock Mini-Cell chamber
• Pre-Cast Nupage 4-12% gel
• 1x MES-TBST Buffer
• CAPS Transfer buffer:
• 200 mL of ethanol
• 4.43 g CAPS
• 0.84 g sodium hydroxide pellets
• Up to 2 L with DI Water
• Owl VEP-2 Electrotransfer Chamber:
• Sandwich + Clip + Sponges
• Immobilin-P transfer membrane (0.2um)
• Whatman paper
• Spatula to crack open pre-cast gel case
• Orbital Shaker
• 5% Milk:
• 100 mL 1XTBST
• 5 g powedered milk
• 5% Milk and 0.02% Sodium Azide:
• 10 mL 5% milk
• 40 uL Sodium Azide
• Primary Antibodies:
• Dilute into sodium azide solution at appopriate concentration
• Can be re-used
• Secondary Antibodies:
• Dilute in 5% milk/TBST solution to appropriate concentration
• Cannot be re-used
• Film Developement Room:
• Red light
• Developer machine
• Saran Wrap
• Film (Do NOT open box in the light)
• Film Cassette
• WesternBright Sirius HRP Substrate Kit:
• WesternBright Sirius™Luminol/enhancer solution
• WesternBright Peroxide Chemiluminescent Detection Reagent
• Scissors

Procedure

Inoculate a Culture of Chlamy:

1. Fill a beaker with 40 mL of TAP.

2. Use a sterile toothpick to smear a few colonies.

3. Wet the side of the beaker by gently sloshing the TAP, and spin the stick against the wet side of the beaker to transfer the colonies into the media.

4. Gently stir and allow to shake at 200 rpm with light until it reaches confluency.

Total Cell Lysate:

5. Transfer 40 mL of cells into a falcon tube.

6. Spin cells down for 5 minutes at 5000 rpm.

7. Resuspend cells in 1 mL wash buffer.

8. Spin the cells again for 5 minutes in a microcentrifuge tube at 4C.

9. Aspirate the supernatant, removing as much as possible.

10. Resuspend cells to around 10^8 cells/mL with fresh 1x PI. Keep on ice.

11. Freeze in liquid nitrogen overnight

12. Thaw samples the next day on ice.

13. BCA ASSAY.

14. Add 1 volume (=volume of cells in 1x PI) of 2x Laemmli gel sample buffer. Keep on ice until all are resuspended.

15. Use pipet tip and then vortex. It will be viscous.

16. Heat at 40C for 30 minutes.

17. Vortex and spin for 5 minutes at full speed in microfuge.

18. Transfer the supernatant to a fresh tube and discard the pellet. Keep on ice until ready to load on a gel.

Western Blot - Gels:

19. Rinse XCell SureLock Mini-Cell chamber with DI water.

20. Rinse pre-cast gel (Nupage 4-12%) with DI water prior to taking it out of the bag.

21. Once pre-cast gel is taken out of the bag, make sure to take the tape off.

22. Place gel into buffer core slot with the front side facing forward.

23. Labeling should be in readable orientation.

24. If not running two gels, place dummy gel or remove slot for second gel.

25. Lock buffer core using gel tension wedge.

26. Fill buffer core with 1X MES-SDS buffer;

Wait for ~30 seconds to make sure that gel box is locked properly and buffer is not leaking. Continue to fill lower chamber until it is half full.

27. Take out comb from pre-cast gel.

28. Wash out wells using p200 (~100 ul).

29. Load ladder using gel-loading 200 uL pipette tips:

Load 7-15 ul of sample using gel-loading 200 uL pipette tips

Do not leave empy wells next to samples: Fill with Laemmli

Only load 7ul of ladder and Laemmli, can load more of sample based on concentration of protein (~7 ug)

Be careful with the final dispensing bubble.

30. Run gel at 120 V *buffer should fill up the core and cover the electrodes outside of the core.

When starting the electrophoresis, ensure that there are no bubbles. Make sure the gel actually starts running.

31. Stop the gel once the samples have moved sufficiently down gel (~45 mins. Less if smaller proteins)

32. While Gel is running:

33. Make 2 L of transfer buffer -

200 mL ethanol

4.43 CAPS

0.84 g sodium

Hydroxide pellets

DI water up to 2L

Mix using a stir bar

34. After Gel is done running: Rinse Owl VEP-2 and sandwich with DI water.

35. Fill Owl VEP-2 with transfer buffer (CAPS).

36. Cut Immobilin-P transfer membrane to appropriate size.

37. Cut Whatman paper to appropriate size.

38. Fill a bin with transfer buffer to soak both sides of the sandwich.

39. Fill a bin with ethanol (large petri dish).

40. When electrophoresis is done, unlock buffer core.

41. Remove pre-cast gel from buffer core.

42. Rinse pre-cast gel with DI water.

43. Use spatula to crack open pre-cast gel case;

Cut off excess gel (bottom of gel, sides, and top).

44. Set Up Sandwich as follows:

i. Lay down Sandwich with BLACK on bottom

ii. sponge

iii. whatman paper:

whatman paper with serological pipet so that there are no bubbles.

iv. Place gel (that has already been cut) so that PROTEINS FACE RED

v. Place transfer membrane on gel, facing proteins. Activate transfer membrane prior to using by soaking it in ethanol for ~10 seconds. Use non-serrated forceps to handle transfer membrane. Make sure it is evenly coated and there are no bubbles (roll out).

vi. Equilibrate transfer membrane with the transfer buffer in the Owl VEP-2.

vii. Whatman paper:

- Roll out whatman paper with serological pipet so that there are no bubbles

- Add some buffer to wet paper

viii. Sponge.

ix. RED on top.

45. Lock Sandwich with plastic clip.

46. Place sandwich in Owl VEP-2:

i. Rubber clip should be facing the top.

ii. Black side of the sandwich should be facing forward when the electrodes are on the right (black side of the sandwich shoudl align with the black electrode, red should face red electrode).

iii. If possible, place sandwich as close to positive electrode.

47. Top off Owl VEP-2 with transfer buffer.

48. Run the electrotransfer at 45 V for 3 hours at 4 C.

49. Remove the sandwich and membrane from sandwich.

50. Equilibrate membrane by placing it in dish with 1X TBST on orbital shaker at 40 rpm for ~5 min.

51. Make 5% milk:

- 100 mL TBST

- 5 g milk

Wester Blot - Blocking and Tagging:

52. Change out 1X TBST with 5% milk/TBST mix and place on orbital shaker at 40 rpm for 30 minutes or overnight (Blocking).

53. Meanwhile membrane is shaking,

i. Make 0.02% (w/v) sodium azide (dilute with 5% milk/TBST) to add to primary antibody to prevent microbial contamination.

ii. Dilute primary antibody and add to sodium azide solution.

54. Rinse membrane with 1X TBST.

55. Add 1X TBST to membrane and place on orbital shaker at 40 rpm for 5 minutes (first wash).

56. Change out 1X TBST and place on orbital shaker at 40 rpm for 5 minutes (second wash).

57. .Change out 1X TBST and place on orbital shaker at 40 rpm for 5 minutes (third wash)

58. Cut membrane with razor blade:

i. Make sure to clean off razor blade with methanol prior to use.

ii. Cut at ladder midline.

59. Place cut membrane in western blot antibody incubating box.

60. Add ~3 mL of primary antibody/sodium azide mix to each slot of box with a sample:

i. Make sure that solution covers entire sample.

ii. Can place saran wrap over slots to prevent spill over.

61. Place antibody incubating box on rocker at 4 C overnight.

62. Change out antibody/sodium azide solution with 1X TBST and place on orbital shaker at 40 rpm for 5 minutes (first wash).

i. Can save antibody.sodium azide solution.

63. .Change out 1X TBST and place on orbital shaker at 40 rpm for 5 minutes (second wash)

64.Change out 1X TBST and place on orbital shaker at 40 rpm for 5 minutes (third wash).

- Make Secondary antibody solution (~5mL, in 5% milk solution).

65. Change out 1X TBST with secondary antibody (diluted with 5% milk/TBST) and place on orbital shaker at 40 rpm for 30 minutes at room temperature.

i. Make sure secondary antibody solution covers entire sample.

66. Change out secondary antibody with 1X TBST and place on orbital shaker at 40 rpm for 5 minutes (first wash).

Western Blot - Film Imaging:

67. Transport samples to film developer room -

BRING:

i. waste container

ii. saran wrap

iii. film

iv. cassette

v. 1.5 mL or 5 mL microcentrifuge tubes

vi. WesternBright Sirius HRP Substrate Kit:

- WesternBright Sirius™Luminol/enhancer solution

- WesternBright Peroxide Chemiluminescent Detection Reagent

vii. p1000

viii. scissors

xi. timer

x. 1X TBST

xi. marker

68. Turn on developer (should take ~15 minutes to be ready to use).

- Wait until red "ready" light illuminates. When ready to use, place used film in developer to wet rollers.

69. Change out 1X TBST and let sit for 5 minutes (second wash).

70. Change out 1X TBST and let sit for 5 minutes (third wash).

71. Prepare enough HRP substrate (1:1) to cover samples.

72. Place membrane on saran wrap and cover samples with HRP substrate for 1 minute.

73. Place membrane on cassette (protein side up).

74. IN THE DARK:

i. Cut film in half.

ii. Cut one corner off of film piece: place this corner in the corner of the cassette.

iii. Place film in cassette:

- do not shift film once placed, place directly on top

iv. Close cassette for appropriate time according to antibody used.

v. Place in developer.

vi. Wait until film is entirely done going through developer.

vii. TURN LIGHTS BACK ON.

75. Mark film:

i. membrane edges

ii. relevant ladder bands

iii. Date

Stripping/Re-Blocking/Re-Tagging:

76. Rinse Membrane 3x in TBST for approx 5 mins each rinse.

77. Cover membrane in small amount of antibody stripping solution.

78. Place on shaker @ 40 rpm for 5 mins .

79. Pour off stripping soluiton.

80. Re-rinse membrane 3 times in TBST for approx 5 mins each rinse.

81. Re-block exposing to 5% milk in TBST solution for 1hr or overnight.

82. Repeat blocking/tagging section of protocol.