LAB HELPER PROTOCOLS

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Autoclaving

1. Prepare items for autoclaving by first covering them in foil.a. Label all items with a small amount of autoclave tape.b. Initial and label all mixtures

2. Place items inside and be sure to line up and shut the door completely, tightening the wheel until locked.

a. The autoclave doors have a double-hinge, so make sure to push in both sides of the door.

3. Sign up on the autoclave sheet and choose a cyclea. DRY: (all items w/o lid or liquid) Press 1 twice.b. WET: (all items w/ liquid) Press 4 twice.

4. After the cycle is complete, check to make sure that the PSI = 0 or mmHg = 0. Open door slightly to release steam.

5. If necessary, allow items to cool 10 minutes before unloading them.a. Use thermal gloves to handle hot items.

NOTES: 6. Dry cycle takes ~45 minutes to complete.7. Wet cycle takes ~1 hour to complete.

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Biohazard Handling

1. Prepare biohazard waste by first knotting the interior bag, followed by knotting the exterior bag.

2. Secure knots with autoclave tape.3. Place bag in autoclave bins (found in autoclave room by waste bin on the counter)

and run through cycle A.4. Replace with two (or one) new autoclave bags found above the refrigerator.

NOTES: 1. Use gloves when dealing with biohazard waste.2. Take autoclaved waste down to the loading dock dumpster.3. Door Code: 268408

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10% SDS

1. Add SDS to beaker with stir bar – carefully.2. (In hood) add ½ of the final ddH2O volume and start to stir slowly.3. Once SDS starts to dissolve, add ddH2O shy of the total amount and stir slowly in

order to avoid foaming of the SDS.

NOTES: 1. Do mixing in the hood.2. Do not breathe in the detergent.3. SDS powder is found in the cabinet by the door.4. Bottle of mixed SDS is found on the shelf next to the hood.5. This protocol will take awhile.

MEASUREMENTS: 1. 10% SDS = 0.1 SDS = 10g SDS / 100 mL ddH2O

a. For 400 mL, use 40g SDS.b. For 500 mL, use 50g SDS.

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Non-HIS Protease Inhibitors

5 mL Package Size: 1. Add 1 mL DMSO, vortex for one minute.2. Add 4 mL of deionized H2O.3. Prepare 200 uL aliquots.

25 mL Package Size:1. Add 5 mL of DMSO, vortex for 1 minute.2. Add 20 mL of deionized H2O.3. Prepare 200 uL aliquots.

STORE @ -20C.

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HIS Protease Inhibitors (Tablets)

1. Tabs are found in the 4C fridge.2. Add 2 mL H2O per tab.

a. Use tabs as needed.3. Nutate at 4C to dissolve.4. Prepare 400 uL aliquots.5. Store at 4C.

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10% APS

1. Mass out 0.5g APS (found in the cabinet) and dilute to 5 mL volume in H2O.a. Vortex and bring up to volume.

2. Aliquot out 50 aliquots at 100 uL per aliquot.3. Place aliquots in the specified freezer box.

a. Mark caps of each tube with a single black dot before placing into the freezer box.

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Cleaning Glass Beads

1. Pour glass beads into a beaker.2. Add bleach and water to the beaker.3. Rinse out and add more bleach.4. Rinse again and add soapy water. 5. Wash thoroughly by rolling the beads in your hand in the beaker.6. Rinse thoroughly with H2O from the tap and add H2O to beads and roll beads

again in hand.7. Repeat this two more times.8. Add DI water and rinse.9. Add ethanol and roll beads around w/ hand in the beaker.10. Place in the hood overnight and allow ethanol to evaporate off.11. The following morning, divide the beads out into about three tall square bottles.12. Keeps the lids loose, keep them secured via autoclave tape.13. Dry autoclave.

NOTES:1. Use gloves.2. Change your gloves between the bleach and H2O step.3. Beads are found in the far left section of the main lab on the 1st bench by the wall.

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4X Lower Gel Buffer

1. Mass out 18.17g Tris base for 100 mL – adjust for amount as needed.2. Pour about 10 mL 10% SDS into a 30 mL medicine cup.3. Measure out 4 mL of 10% SDS per 100 mL final solution volume and add the

Tris-base.4. Bring up to just shy of the final volume, but enough to cover the pH meter. 5. Cover the beaker, label it, and cool for a couple of hours.6. pH close to 8.8 using 6.0N HCl.7. Bring the volume up to the final volume.8. Adjust the pH as necessary.

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Making Destain Solution

1. In a beaker, combine 10% MeOH, 10% HOAc, and 80% H2O.2. Mix in a hood and add to appropriate bottle in hood.

NOTES:1. First add ½ the water, then the acid and methanol, then bring up to the final

volume with water.2. For 4L: use 400 mL methanol, 400 mL acetic acid, and 3200 mL water.3. If there is an excess, dispose of it into the coomassie waste bottle.

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Cleaning Pipettes

1. Connect the hose to the faucet and run hot water through the pipette container at the same rate that the water is flowing out of the bottle of the container.

2. Add 2 ½ tbsp of detergent.3. Run hot water for 30 minutes.4. Fill with DI water and repeat drain/fill cycle for another 30 minutes.5. Lay down paper towels to cover the cart.6. Let pipettes dry on cart with paper towels.

NOTES:1. Wear gloves.2. Immerse pipettes in H2O.3. If floods, refill and attempt to reach a steady state again.

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Coomassie Dye

1. Mix together (for 1L solution)a. 1g coomassie G-250 (0.1%)b. 400 mL methanol (40%)c. 100 mL acetic acid (10%)d. Rest water 1L

2. Stir on stir plate in the hood.3. Prepare gravitational filter system for each bottle to be filled.4. While filtering, be sure to keep mixture stirring.5. Date and cap the bottle in the hood.

NOTES:1. Wear a coat and gloves.2. Takes about 2 hrs to gravity filter 1L.3. Place paper towels around area to sop up spilled coomassie.4. Coomassie is a stain that will ruin clothes, stain skin, etc; be careful, yo.

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Cleaning NMR Tubes

First Day1. Fill NMR tubes 1/3 fill with nitric acid using an NMR pipette. Allow to sit

overnight.

Second Day2. Use NMR pipette and empty NMR tubes, putting waste in Tris solution.3. Clean tubes by connecting apparatus to a vacuum system.4. Cap the bottom of the NMR tube and place into the slot…5. Squirt water, then cuvette cleaner, then rinse with ddH2O, then 1 squirt of bleach,

then more H2O, and finally a shot of acetone.6. Repeat.

NOTES: 1. Wear gloves, overcoat, and goggles.2. Any excess nitric acid can go into the 1M Tris-base jar in the acids cabinet. The

indicator is yellow when solution is too acidic. Add tris-base until blue again.a. To make this buffer with indictor, make about 250 mL 1 M solution and

add about 25 mg bromophenol blue.

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1000X Vitamins

1. Measure out 50 mg of each of the following vitamins.a. Biotin -FRIDGEb. D-Pantothenic Acid - FRIDGEc. Biz. – Cyanocobalamine d. Choline Cl – ROOM TEMPe. Folic Acid – ROOM TEMPf. Niacinamide – ROOM TEMPg. Riboflavin – ROOM TEMPh. Pyrodoxial HCl – FREEZER

2. Mix together and bring to a final volume of 50 mL in a covered conical vial; wrap in foil.

3. Let stir until all vitamins have gone into solution; final color should be a red/orange.

4. Sterile filter into another conical via syringe filtering.5. Aliquot out 1 mL/aliquot (should get about 50 aliquots).6. Put into box in freezer.

NOTES: 1. Mass out room temperature vitamins first. 2. Wrap conical vial in foil, substances are light sensitive.

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10X TBS

1. Mix together:a. 292.2g NaClb. 24.22g Tris-basec. Bring to final volume with 1 L H2O

2. pH to 7.5

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1M DTT

1. Mass out 1.54g DTT and add 10 mL dH2O in a 15 mL conical.2. Vortex3. Make 200 uL aliquots.4. Mark tops of microcentrifuge tubes with a black X.

NOTES:1. Do in hood.2. Should make ~55 aliquots.

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Western Blot Paper

1. Sheets are found in to chromatography room in the cabinet under ‘used fraction tubes.’

2. Using a piece from the Western drawer as a template, use the paper cutter in the dry lab.

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Antibiotic Plates

1. Mix together:a. 10g NaClb. 5g yeast extractc. 10g tryptoned. 15g agare. 1L ddH2O

2. Autoclave in a fernbach flask on cycle 4 w/ a magnetic stir bar in the flask!3. Add 1 aliquot/L of the appropriate antibiotic stock to the flask once the

temperature is less than or equal to 45C degrees.a. Adding the antibiotic while the media is still extremely hot will inactivate

the antibiotic.4. Plate media and let solidify.5. Label plate with a sharpie.

a. Kanamycin – Blueb. Ampicillin - Red

NOTES:1. If making 2L, cool one on bench top and the other in an ice bath w/ a stir plate.2. Add antibiotic once cooled.3. Hold bottom of flask to wrist to check for coolness (

a. should be about as warm as a baby bottle – AKA: “don’t give the baby scalding-hot fluids”

4. Move plates carefully when the agar is settling.

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Electrocompetant Cells

Dry Autoclave:1. 2 large centrifuge bottles w/ caps.2. 2 small centrifuge bottles w/ caps. (SS-34)3. 1 glass jar of 1.7 mL epi tubes.4. 1000 uL and 200 uL tips.5. 10 mL and 25 mL pipettes.6. 150 mL Erlynmeyer flask

Wet Autoclave: 1. 2 1L flasks of LB2. 2 1L bottles of water

a. If needed, 1L LB for starter culture (only need 50 mLs).

Prepare: 3. 10% glycerol (sterile filtered)

Night Before (anytime after 4pm):Starter Cultures

4. Inoculate ~50 mLs of sterile LB w/ cells from store stock in 150 mL EF (autoclaved)

a. Inoculate by taking a small swipe off with a sterile pipette tip and swirl around in the LB

b. Use can also pipette up and down.5. Label EF with type of cell, date, and initials.6. Grow at 37C and 250 rpm in warm room.

Day Of: *set shaker to 4C

1. Inoculate each liter of LB with 10 mL of starter culture (be sure to keep sterile).2. Grow cells at 37C and 250 rpm until you reach optical density.3. Meanwhile, put all sterile flasks of water and centrifuge tubes in the cold room.4. Check OD after about 2 hours, then at 15 minute intervals.

a. OD desired: 0.6 – 0.95. When OD is reached, transfer the LB sterily to two sterile large centrifuge bottles.

a. DO NOT fill bottles to the top; they will leak in the spin cycle.b. Balance with two other large centrifuge bottles.

6. Spin using 4000 rotor.a. Set rotor number to 33b. 10 minutes @ 4200 rpms

7. When cycle is complete, decant each bottle and resuspend in 500 mLs of sterile H2O.

8. Place in 4C shaker to resuspend.9. Balance out and respin for 10 more minutes.

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10. Decant again and resuspend in 250 mL each bottle sterile H2Oa. Resuspend in 4C shakerb. When resuspended, combine into 1 bottle for total of 500 mLs.c. Balance out and spin again for 15 minutes.

11. Decant and resuspend in 20 mLs of sterile 10% glycerol.12. Once resuspended, transfer to one small centrifuge tube (SS-34 tube)13. Balance out and spin down at 5000 rpm for 15 minutes.14. Get liquid nitrogen while spinning.15. Decant and resuspend in 2 mLs sterile…

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