Autoclaving

AUTOCLAVES:
2 department autoclaves found in 811 Riffe Building

SUGGESTED SETTINGS:
Liquids Setting Temp Time Setting Approximate run time
Small Volumes 1270C 30 minutes sterilization 50-55 minutes
Large Volumes 1270C 45 minutes sterilization 1 hour, 10 minutes
Wrapped Setting
1210C 45 minutes sterilization 1 hour, 45 minutes
30 minutes drying time

DON'T FORGET:
Place autoclave tape on materials
Liquids should fill bottles approximately half way
Bring autoclave gloves with you
If you change the settings to non-standard values change them back

For more, see Long Form.

Disposal

Disposing of Burn Boxes

Box should weigh no more than 30 pounds (1/3-1/2 full)
Close and tape securely with packing tape
Label with lab and room number
Fill out form at www.ehs.ohio-state.edu/env-affairs/biohazard.html
Place box in the hallway for pickup

For information on disposal of other lab reagents and supplies see the Long Form.

Using DNA Strider 1.2

This program allows for manipulation of sequence data.

Sequence can be copied and pasted, translated, checked for enzyme sites, displayed in various formats and searched for specific sub-sequences.

Strider has a sequence length limit of 20 kb (20,000 bases). To work with larger sequence copy and paste it in two parts into two files.

For more information, see the Long Form.

Getting Reagents On Site

Enzymes and other Reagents from the Dean Lab

See the Long Form.

Enzymes from the Fermentas Freezer

1. Get the freezer card from the office supply drawer. You can find the pin number in the front of the Enzyme notebook.
2. Take the card and some ice with you to Room 968.
3. Follow the instructions on the freezer to obtain your enzyme or other reagent. Put in ice immediately. Remember to put enzymes directly in the ­200C freezer and add the appropriate information to the enzyme freezer sheet.
4. Record the new card balance on the Post-it Note on the card.

For a list of freezer contents, see the Long Form.

Qiagen Products from the Ostrowski Lab

1. The Ostrowski Lab maintains a Qiagen cabinet. They accept 100-W forms as payment.

For more, see the Long Form.

Mailing Parcels

1. Properly secure your parcel, package, etc. (Envelopes and such can be found in the bottom drawer directly across from the filing cabinet in lab).
2. Make sure the envelope or box has our return address and the address you are mailing to. (Tape down white scraps of paper with addresses written or typed on)
3. Take the parcel to the Department office (Room 984).
4. As you enter, on your immediate right you will see stamps hanging from a rotating wheel. You will find the ink pad in the top drawer nearest the wheel.
5. Select the stamp that reads "0340-200340-36I". Dip this stamp in the ink and then press the stamp on your parcel once below the return address. (This stamp identifies our department)
6. If your parcel needs to be sent overnight, select the stamp that reads "DHL Worldwide Express". Dip this stamp in the ink and then press it on your parcel two or three times in various places.
7. If you are not in a rush you can simply place your parcel in the outgoing mail box in the Mail Room (907). In this case, the parcel will go downstairs the next day and be sent in the mail. If you are in a rush and are sending it "DHL Worldwise Express" you can make sure it goes out the same day by placing it in the DHL drop box near Lincoln Tower.

Making a Worm Pick

1. Get the following:
- platinum wire
- 1 glass pasteur pipet
- paper towel
- forceps
- scissors
- gas or Bunsen burner
2. Place pipet in paper towel to prevent escape of broken glass
3. Use forceps to break pipet near where it begins to narrow.
4. Using scissors cut off a piece of platinum wire about 1.0-1.5 inches (3.5-4.0 cm) long.
5. Light burner.
6. Holding platinum wire piece with forceps, insert it about 1.0 cm into the broken tip of the pasteur pipet.
7. Hold the tip of the glass (with wire also held in via forceps) over the flame until the glass begins to melt.
8. Use the forceps to press and sculpt the glass over the platinum wire. Repeat until the platinum wire is secure.
9. Sculpt the tip of your wire as desired for picking worms.
10. Clean up/dispose properly.

For more, see the Long Form.

Ordering

Ordering Primers

1. After you have designed your primers fill out a purchase order form including the total price ($0.35/base).
2. Place the form in Marsha's Box and write a note requesting a P.O. number so you can order on-line.
3. When you receive the P.O. #, order primers at http://www.mwgbiotech.com/services/orders/head.htm
4. Double check primer sequences before submitting order.
5. Place the order sheet in the blue orders received folder.
6. Primers will be shipped to us in 1-2 business days.
7. When they arrive follow Receiving Primers and Making Working Stocks under the protocol (PCR Polymerase Chain Reaction).

For more, see the Long Form.

Sequencing (Neurobiotechnology Center)

Prepare DNA and primers to standards.
Fill out an order sheet for the reactions with all information. Also fill out a 100-W form for the appropriate amount ($15.00/rxn).
Take the samples to the Neurobiotechnology Center and leave them in the fridge in the lobby. Leave the forms in the box there.
You should receive results in two-four working days.

For more, see the Long Form.
Sterilizing

Glassware: Washing Glassware and Using the Glassware Facility

Washing Glassware

Non-contaminated labware should be rinsed with hot water, left to dry, and once dry placed on the glassware cart.
When the glassware cart is filled it is taken to the Glassware Facility (2nd Floor). It can be picked up 1 or more days later.

The Glassware Facility does not accept the following: Graduated Cylinders larger than 500 ml, Funnels with stems smaller than 1 cm diameter, Syringes, Watch glasses, Carboys, Staining containers, Stirring rods, spatualas, and stir bars. These items should be washed in lab, dried and returned to their appropriate place. If necessary autoclave sterilization is an option for some of these items.

For special cases ­ contaminated labware, caps, culture tubes with media and bacteria see the Long Form.

For more on the Glassware Facility, see the Long Form.

Sterilizing Centrifuge Tubes

1. Tubes and lids should be clean before autoclaving
2. Use foil to wrap large lids completely and to cover the top of tubes. Smaller tubes should be autoclaved with lids on.
3. Apply autoclave tape.
4. Autoclave Wrapped setting (45 min/30 min).

For more, see the Long Form.

Sterilizing Double Distilled Water

1. Add double distilled water to a 50 ml plastic tube.
2. Using a P1000, dispense water in 1.0 ml aliquots to sterile 1.5ml tubes (with flexible tops).
3. Place tubes in a beaker and cover beaker with a foil lid.
4. Apply autoclave tape to foil.
5. Autoclave Wrapped setting (45 min/30 min).

For more, see the Long Form.

Sterilizing Glass Pipets

1. Transfer about 100 pipets from storage canisters to the Nalgene pipet washer.
2. Hook up tube to water nozzle and run hot water through cleaner for 10 or more minutes.
3. Turn off water.
4. Disconnect tubing.
5. Let the pipets dry overnight.
6. Place pipets in metal canisters.
7. Secure lids with autoclave tape.
8. Autoclave Wrapped setting (45 min/30 min).

For more, see Long Form.

Sterilizing Plastic Tubes

1. Fill a beaker with plastic tubes.
2. Cover beaker with a foil.
3. Affix autoclave tape.
4. Autoclave Wrapped setting (45 min/30 min).

For more, see the Long Form.

Sterilizing Glass Test Tubes

1. Only washed or new glass test tubes should be autoclaved by sterilization.
2. In a white test tube rack (72 tube capacity) place clean or new tubes with plastic caps one.
3. Wrap a small piece of autoclave tape around the rack (this will indicate that all tubes in the rack are sterile).
4. Autoclave Wrapped setting (45 min/30 min).

For more, see the Long Form.

Sterilizing Tips

1. Put on gloves for handling tips.
2. White tips and blue tips must be added by hand to empty racks. Yellow tips are added via a quick dispenser which is easy to use.
3. Once tips are added, autoclave tape should be applied at the closure of each rack.
4. Autoclave Wrapped setting (45 min/30 min).

For more, see the Long Form.

Sterilizing Toothpicks

1. Wearing gloves fill a clean or new glass test tube with toothpicks (about 40-50 toothpicks in two layers).
2. Place a plastic test tube cap firmly but not tightly.
3. Use a small piece of tape to secure the lid to the tube.
4. Autoclave Wrapped setting (45 min/30 min).

For more, see the Long Form.

Streaking Bacterial Plates

1. Obtain plates of the appropriate type (usually LB).
2. Label bottom of each plate with bacterial strain name, your initials, and the date.
3. Prepare area with:
- gas burner
- platinum innoculating loop
- bacterial source (liquid, plate or other)
- plates
4. Flame loop.
5. Touch the loop to the innoculum bateria (liquid or colony) and move the loop in a smooth zig zag across the surface of the first quarter of the plate.
6. Flame loop.
7. Streak again, starting from the end of the last streak and touching it only once. Streak out another quarter of the plate surface.
8. Repeat steps as above for the rest of the plate.
9. Incubate plates at the proper temperature (usually 370C) for many hours. 10-16 hours is suggested for many bacteria.
10. Clean up.

For more, see the Long Form.

Worm Plates
Plates without worms can be stored at RT or at lower temperatures (40C) to increase the survival of the plates. Plates are spread with bacteria (Op-50 strain of E. Coli) to serve as nutrition for worms.

Plates containing worms can be stored in either the 150C or 200C incubator. The worms will adjust to whatever temperature they are stored at. They eat more bacteria and grow and reproduce faster at 200C.

Removing Bacterial Contaminants from Worm Plates

Method 1. Transfer a single hermaphrodite from the contaminated plate to a new, non-contaminated plate. Wait for another generation to grow up (hopefully uncontaminated).

Method 2. Transfer a single hermaphrodite from the contaminated plate to a new, non-contaminated plate. Wait 1 hour. Transfer this worm again to a new, non-contaminated plate. Wait 1 hour. Transfer the worm one last time to a new, non-contaminated plate. Wait for another generation to grow up (hopefully uncontaminated).

Method 3. Apply a few milliliters of solution to the contaminated plate from a squirt bottle of 10% Bleach solution. Swish the solution around. Wait a few minutes and then pour off carefully. The bleach will kill all the worms and dissolve their bodies, however the fertilized eggs will survive to produce worms. As soon as these worms develop, transfer them to a new, non-contaminated plate.

For more on Worm Plates, see related protocols.

General Lab Tips

Emergencies

Emergency Numbers: 911 or 3-8333 (University Hospitals Emergency Room)

Eye Wash: Located on the right side of the sink nearest the door to 941. To turn on squeeze the handle. To turn off pull on the piece that sticks out from the base of the handle.

Chemical Shower: Two showers are found on the floor. One outside 956 (Chang Lab). The other is outside 916 on the opposite side of the floor. Both are roughly equidistant from the lab.

Sterile (Aseptic) Technique

Sterile technique is most important for sensitive applications involving living organisms like making, pouring and spreading plates, preparing cultures, using LB, transferring worms, etc. These techniques are not always necessary for successful experiments, but they are good habits to form.
1. Use 70% Ethanol (squeeze bottle) to wipe down lab surfaces before sensitive applications
2. Use sterile solutions and materials for applications. If you are not sure something is sterile then do not use it as if it is
3. Set up your working area to minimize the amount of hand and arm movement you have to make
4. Have everything ready ahead of time so that you do not have to scramble to find something
5. Wear gloves when appropriate for your own protection and to avoid contamination of samples and equipment
6. Open bottles at a 450 angle to minimize air-born contamination
7. If you put down a cap or lid, put it face down to lessen the chance of air-born contamination
8. Always flame glass pipets and open bottles before use. You can also flame bottles again before closure
9. Pipet gently and do not swirl (this minimizes aerosols)
10. Do not leave bottles open longer than they need to be
11. Stop what you are doing when any of the following happen:
- You pipet too far up the pipet (and potentially contaminate the pipettor). Stop and clean the pipettor with 70% ethanol at the least. Change any filters if necessary.
- Touching the tip of the pipet against a bottle, the bench or anything solid. Discard the pipet.
- Dropping an opened container or tube to the ground. Discard it.
- Reusing pipets. Do not reuse pipets for different applications. They are more likely to pick up contaminants.

Using Equipment

If there is a sign up sheet or book, then sign up. Sign up even if you are sure no one else will be using; you never know when things will change. This also provides a record of who has used the equipment.
Follow all guidelines for the equipment. Turn it off when you are done if appropriate.
If there is a problem that you can not deal with make sure to find and tell someone who can. You may also want to leave a note on the equipment indicating any problems you had.

Pipetting

For more on pipetting and using specific types of pipets in our lab (glass, disposable, etc.), see the Long Form.

Lab Protocols

Making Competent E. Coli Cells
(adapted from Dave Deitlerer's protocol)

1. Innoculate 6 ml of culture from stock and grow overnight.
2. Dilute 1:100 in LB (6 ml > 600 ml) in a 2.0 L flask.
3. Grow in shaker-incubator approximately 2 hours.
4. To determine when the cells are in the exponential growth phase take optical density readings on samples. The target OD is about 0.3 at 600nm.
5. Place 4 sterile, 150ml centrifuge tubes on ice.
6. Weigh the tubes and adjust with aliquoted LB until they are closely balanced.
7. Chill on ice 5 minutes.
8. Centrifuge 3X1,000 RPM for 5 minutes.
9. Pour off the supernatant into liquid waste flask.
10. Resuspend each pellet in 100 ml of Ca/Glycerol buffer. Vortex to resuspend.
11. Balance the tubes by weight using Ca/Glycerol.
12. Centrifuge at 3,000 rpm for 5 minutes.
13. Repeat steps 9-11.
14. Chill on ice for 30 minutes.
15. At this time obtain enough dry ice to cover the bottom of an ice container.
16. Centrifuge at 3,000 rpm for 5 minutes.
17. Pour off the supernatant into liquid waste flask.
18. Resuspend each pellet gently in 12.5 ml (TE?).
19. Place approximately 100 sterile, 1.5 ml tubes on a bed of dry ice to chill.
20. Using a P1000 aliquot resuspended cells, 0.5 ml to each tube.
21. Store tubes at ­700C for long term. To use, thaw on ice and use immediately.
22. Make sure to check the competency of the batch of cells, by performing a transformation as detailed below.

For more details, see the Long Form.

Transformation of Competent E. Coli cells
(adapted from Akira Shimuzu's protocol)

1. Thaw source BS DNA (1 ng/ul).
2. Thaw 1 tube of competent cells.
3. While reagents thaw, set up 2 transformations in sterile, 1.5 ml tubes (for example):
Bluescript DNA Negative Control
20 ul 10X TF Mix 20 ul 10X TF Mix
1 ul bluescript (1 ng/ul) 80 ul sterile water .
79 ul sterile water
4. Proceed immediately to add 100 ul of competent cells to each tranformation.
5. Tap to mix and ice transformations for at least 20 minutes.
6. Place transformations at RT no more than 10 minutes.
7. Add 1.0 ml LB to each transformation.
8. Shake/Incubate transformation for 45 minutes to 1 hour.
9. Setup LB and LB carb plates and serial dilution tubes for later use:
- 1 LB plate labeled 'Negative Control'
- 1 LB Carb plate labeled '1/6 ng/ml BS'
- Label 4, 0.6 ml tubes 1/60, 1/600, 1/6,000, 1/60,000 (respectively) and add 180 ul LB to each
- Label 4 LB Carb plates 1/60, 1/600, 1/6,000, 1/60,000 (respectively)
10. Get:
- Glass bacterial spreader
- tub of ethanol
- Bunsen burner
- Turn table for spreading
11. Retrieve transformations after at least 45 minutes.
12. Perform serial dilution of BS transformation (20 ul added to each subsequent dilution).
13. Add transformations/serial dilutions to plates and spread using sterile technique:
- 200 ul of 'negative control' to LB plate
- 200 ul of 'BS' to LB Carb Plate labeled 1/6 ng/ul
- 180 ul of each dilution to their appropriately labeled LB Carb plates
14. Incubate overnight (at least 16 hours) and check for cell competency as below.

For more, see the Long Form.

Checking Cell Competency

Expected plated results of transformations if cells are competent:
- LB negative control plate has no colonies
- 1/6 ng/ml ­ very many colonies
- 1/60 ng/ml ­ many colonies (too many to reasonably count)
- 1/600 ng/ml ­ some colonies, countable
- 1/6,000 ng/ml ­ few colonies
- 1/60,000 ng/ml ­ very few or no colonies
Count the colonies on the plate which is easy to count. This count is the number of colony forming units (cfus) for that plate. Multiple the cfu by the inverse of the concentration of that plate. Then multiple this amount by 1,000 since there are 1,000ng/1ug. This will give you a final number in the units (cfu/ug). If this number is between 1 X 107 and 1 X 108 than it is acceptable and you have competent cells.

Example:

40 colonies on the 1/600 ng/ml plate
multiplied by 600 = 24,000
multiplied by 1,000 = 24,000,000 cfu/ug

This is within the acceptable range and the cells are judged to be competent.

Receiving Cosmids

Cosmids will come via mail in an envelope. The tubes contain a "stab" where you can observe bacterial growth. This is the area from which you want to inoculate. Upon receiving cosmids you want to perform 2 protocols:

Make a freezing stock

Label a freezer tube for each cosmid.
Get the following: test tube containing sterile toothpicks, forceps, burner.
To each tube add 50 ul of LB medium and 50 ul of 30% glycerol.
Light burner.
Open the tube that contains toothpicks.
Flame open tube mouth and forceps.
Use forceps to grab one toothpick.
Flame the tube mouth and place the lid back on.
Direct toothpick so that it penetrates the stab of the desired cosmid.
Immerse toothpick in the matching freezer tube to inoculate.
Discard toothpick into sharps box and close the lid of the freezer tube.
Repeat steps 5-11 for all cosmids.
Place tubes in the appropriate box at ­70 C.
Do not forget to enter stock information into the data base, and the Strain, Freezer Box Map, and Plasmid/Cosmid Log notebooks. For more information on how to do any or all of these things , see Making Freezing Stocks.

Streak out plates, Mini prep, and Analyze

1. Get an LB/antibiotic plate for each cosmid (NOTE: make sure the plate matches the antibiotic resistance for the cosmid).
2. Label the plate (base) with the cosmid strain name, date and your initials.
3. Get the following: test tube w/ sterile toothpicks, forceps, burner.
4. Light burner.
5. Open the tube that contains toothpicks.
6. Flame open tube mouth and forceps.
7. Use forceps to grab one toothpick.
8. Flame the tube mouth and place the lid back on.
9. Using the toothpick, streak out the cosmid on the appropriate plate using the standard technique (For more, see Streaking Plates).
10. Discard toothpick into sharps box and close the lid of the freezer tube.
11. Repeat steps 5-10 for all cosmids.
12. Place the plates upside down (lids down) in the 370C incubator. Let them grow for 12 hours or more.
13. The next day pick colonies and use them to inoculate liquid cultures in LB solution containing the appropriate antibiotic. Avoid large "blobby" colonies, and instead pick smallish (but not pinprick) colonies. It is standard to grow at least 2 cultures per cosmid. (See Making Liquid Cultures for more information on this process)
14. Using the liquid cultures, make alkaline mini preps for each cosmid. (See Mini Prep (Alkaline Lysis) for more information)
15. Set up appropriate digests for your cosmid preps. Assay the cosmid recovery by running digests on a DNA gel.

DIG (non-radioactive) labeling of DNA

1. This protocol is adapted from page 10 of DIG High Prime DNA Labeling and Detection Starter Kit II (Roche Molecular Biochemicals ­ Cat. No. 1 585 614). Use this as an additional resource.
2. Thaw template DNA on ice if required.
3. Fill a 2000 ml beaker with about 700 ml of water.
4. Place the beaker on stirrer/hot plate.
5. Turn the heat knob to 8. It will take about 10 minutes for the water to come to a boil.
6. In a small, sterile, labeled tube make a 16ul reaction containing:
1 ug template DNA (DNA must be quantified in order to determine volume to add)
sterile ddH2O to bring reaction volume to 16 ul
(Example ­ 100 ng/ul template. Reaction would be 10 ul template and 6 ul sterile ddH2O)
5. DNA Denaturation Step: Float the reaction tube in the boiling water for 10 minutes.
6. With about 3 minutes left, make an ice and water mixture in a styrofoam container. Get DIG-High Prime (Vial 1) and put on ice.
7. When the 10 minutes is up quickly submerge the float in ice water.
8. Tap the DIG-High Prime to mix. Add 4.0 ul to the denatured DNA. Tap to mix. Briefly spin down.
9. Put the DIG-High Prime back in the enzyme freezer.
10. Incubate the reaction at 370C for 1 ­ 20 hours. (Increased time results in increased labeling in an approximately linear relationship)
11. Stop the reaction by adding 2 ul of 0.2M EDTA (pH 8.0) AND/OR by heating to 650C for 10 minutes.
12. Continue the hybridization process by following the protocol for Semi-quantitative determination of labeling efficiency AND/OR DNA Fixation.

Semi-quantitative determination of labeling efficiency

Theory: Probe is applied to membrane along with control DNA. The two are compared to determine the labeling efficiency of your sample.

This protocol is adapted from pages 12-14 of DIG High Prime DNA Labeling and Detection Starter Kit II (Roche Molecular Biochemicals ­ Cat. No. 1 585 614). Use this as an additional resource.

The following steps cover the recipes for necessary reagents:

Making Washing Buffer (1 L)

1. Fill a 1 L flask with about 800 ml.
2. Add 11.61 g of Maleic Acid (0.1M).
3. Add 8.76 g of Sodium Chloride (0.15M).
4. Adjust pH to 7.50 (at 200C) by adding NaOH pellets:
- Initial pH will be about 1.25
~ 40 NaOH pellets adjusts to pH of 7.50
Add smaller amounts as you near the 7.50 mark
With this solution it is easy to overshoot! (so be careful, slow and accurate is better)
Use drops of 1M HCl to bring the pH down (1 drop will lower the pH ~ 0.01 units)
5. Using a glass pipet, add 3.0 ml of Tween 20 (0.3% v/v) to the flask.
6. Pour the solution into a 1000 ml graduated cylinder.
7. Using dH2O bring liquid level to 1000 ml.
8. Pour the solution into a sterile, 1.0 L screw top bottle.
9. Label bottle. (This solution does note need to be sterilized and is stable at RT)

Making Maleic Acid Buffer (1 L)

1. Fill a 1 L flask with about 800 ml.
2. Add 11.61 g of Maleic Acid (0.1M).
3. Add 8.76 g of Sodium Chloride (0.15M).
4. Adjust pH to 7.50 (at 200C) by adding NaOH pellets:
- Initial pH will be about 1.25
~ 40 NaOH pellets adjusts to pH of 7.50
Add smaller amounts as you near the 7.50 mark
With this solution it is easy to overshoot! (so be careful, slow and accurate is better)
Use drops of 1M HCl to bring the pH down (1 drop will lower the pH ~ 0.01 units)
5. Pour the solution into a 1000 ml graduated cylinder.
6. Using dH2O bring the liquid level up to 1000 ml.
7. Pour the solution into a sterile, 1.0 L screw top bottle.
8. Label bottle. (This solution does note need to be sterilized and is stable at RT)

Making Detection Buffer (1 L)

1. Fill a 1 L flask with about 800 ml.
2. Add 12.11 g of Tris (0.1M).
3. Add 5.84 g of Sodium Chloride (0.1M).
4. At this point the pH must be adjusted to 9.50 (at 200C) by adding concentrated HCl:
Wear gloves, lab coat, and goggles while working with HCl since it is very caustic
Initial pH will be about 10.60
You will need to add ~ 500 ul of conc. HCl to achieve a pH of 9.50
5. Pour the solution into a 1000 ml graduated cylinder.
6. Using dH2O bring the liquid level up to 1000 ml.
7. Pour the solution into a sterile, 1.0 L screw top bottle.
8. Label bottle. (This solution does note need to be sterilized and is stable at RT)

1X Blocking Solution (2 X 10 ml)

1. This solution must always be made fresh on the day it will be used.
2. Thaw 2, 1.5 ml tubes of 10X stock at RT (If stock is unavailable see Long Form on how to make more).
3. Add 9.0 ml maleic acid buffer to each of 2, sterile 15ml tubes.
4. Make1X working solutions by adding 1.0 ml of thawed 10X blocking solution to each tube and mixing.
5. Label one tube as 1X Blocking Solution and one tube as Antibody Solution. The blocking solution is now ready to use. The Antibody Solution needs antibody to be added (see below).

Antibody Solution (10 ml)

1. Customarily this solution will be made fresh each time (unless you recently performed the protocol since this solution only stores up to 12 hours). If making fresh, make only after fresh 1X Blocking solution has been prepared.
2. Balance and centrifuge the anti-digoxigenin-AP (vial 4, stored at 4C) at 10,000 rpm for 5 minutes.
3. After spin, take 1.0 ul of anti-digoxigenin-AP from the surface and add it to your labeled tube of 1X blocking solution (1:10,000 dilution).
4. Mix thoroughly. The solution is ready to be used.

Main Protocol

1. This protocol is adapted from pages 12-14 of DIG High Prime DNA Labeling and Detection Starter Kit II (Roche Molecular Biochemicals ­ Cat. No. 1 585 614). Use this as an additional resource.
2. Obtain wet ice.
3. Thaw control DNA dilutions (tubes #2-9) on ice (stored at ­20C). Also thaw your sample probe DNA.
4. You will need to make up a similar dilution series (tubes #2-8) for your sample probe. The dilution series should be made up according to the following chart:

Tube DNA (ul) From tube # DNA Dilution Buffer (vial 3) (ul)
1 - original -
2 5 1 495
3 15 2 35
4 5 2 45
5 5 3 45
6 5 4 45
7 5 5 45
8 5 6 45

5. If not already done, make 10 ml of 1X Blocking solution and 10 ml of Antibody solution.
6. Using gloves place a small strip of nylon membrane on a paper towel.
7. Cut a piece from the upper right corner of membrane to indicate orientation.
8. Peel back the membrane cover from the top of the membrane. Dispose of the cover.
9. Add 1 ul spots of control dilutions (tubes #2-9) to the membrane in a logical row.
10. Add 1 ul spots of sample dilutions (tubes #2-8 + control tube #9 if desired) to the membrane in another row paralleling the first.
11. Take off the back cover from the membrane and dispose of the cover.
12. UV Crosslink the membrane.
13. Transfer the membrane to a sterile, 50 ml plastic tube. Use a glass pipet to add 20.0 ml Maleic Acid Buffer.
14. Place the tube on a nutator at RT. Turn the nutator on and make sure the solution washes the whole membrane.
15. Let the membrane shake for 2 minutes. Pour off.
16. Add 10.0 ml Blocking Solution.
17. Shake at RT (nutator) for 30 minutes. Pour off.
18. Add 10.0 ml Antibody Solution.
19. Shake at RT (nutator) for 30 minutes. Pour off.
20. Add 10.0 ml Washing Buffer.
21. Shake at RT (nutator) for 15 minutes. Pour off.
22. Add 10.0 ml Washing Buffer.
23. Shake at RT (nutator) for 15 minutes. Pour off.
24. Add 10.0 ml Detection Buffer.
25. Shake at RT (nutator) for 2-5 minutes. Pour off.
26. Using blunt force tweezers place the membrane with DNA side facing up in a plastic development folder.
27. Add 4 drops (~100 ul) of CSPD ready-to-use (light-sensitive, stored at 4C).
28. Immediately cover the membrane with the top sheet of the folder and spread the substrate evenly and without air bubbles over the membrane.
29. Let the folder incubate for 5 minutes at RT.
30. Seal the edges of the development folder with tape.
31. Expose the folder to X-ray film for 20 minutes (see Exposing Film).
32. Develop the film (see Developing Film).
33. Look at the film and compare the intensity of your sample dilutions with the control dilutions. This indicates the semi-quantitative efficiency of the labeling.

Hybridization

Preparation of DIG Easy Hyb Solution from Kit (done once when kit is received)

1. Turn on the water bath and heat to 370C.
2. Wear gloves since this solution is caustic.
3. Remove the bottle of DIG Easy Hyb Granules (bottle 7) from the kit.
4. Measure 64.0 ml of sterile ddH2O in a sterile, 100 ml graduated cylinder.
5. Unscrew the bottle and hold it in the 370C bath. Add about half (32.0 ml) of the water to the bottle.
6. Use a spatula to stir the solution as you hold the bottle in the bath.
7. When the granules begin to go into solution well (1-2 minutes), add the rest of the water and continue stirring.
8. When the granules are completely in solution (an additional 2 minutes or so) it should appear clear.
9. The solution is now ready to use.
10. Store at 40C when not in use. Take aliquots as needed for protocols. These aliquots can be retained and reused if they are stored at 40C.

Making 20 X SSC (used to make subsequent dilutions)

1. Fill a 1.0 L flask with about 700 ml dH2O.
2. Add 175.3g of NaCl.
3. Add 88.2 g of Sodium Citrate.
4. Adjust pH to 7.00:
Initial pH will be about 7.25
Add about 15 drops of 1M HCl to bring the pH close to 7.00
5. Pour the solution into a clean, 1000 ml graduated cylinder.
6. Add dH2O to bring the liquid level to 1000 ml.
7. Dispense the 500 ml solution each into two 1000 ml screw top bottles.
8. Autoclave (Liquids Setting, 1270C, 30 minutes). Retrieve after 50 minutes.
9. Label bottles. The solution may now be used or diluted as needed.

Pre-Hybridization

(NOTE: Pre-hybridization and hybridization temperatures may vary depending on the nature of your experiment. Successful Cross-species hybridization usually requires a lower temperature)
Thaw DIG Easy Hyb (without probe) at RT or in a water bath set to 550C.
Turn on the Robbins Model 1000 Hybridization Incubator or other Hyb/Incubator.
Set Hybridization Incubator to 550C.
Add 10.0 ml of DIG Easy Hyb to a clean hybridization tube (35X300 mm).
Place the tube and an empty balance tube opposite each other in the Hyb/Incubator.
Wait until the Hyb/Incubator and its contents are heated to 550C.
Add your membrane/filter to the tube with DIG Easy Hyb for pre-hybridization (for tips see the Long Form). Make sure the solution washes the whole membrane/filter.
Let the membrane pre-hybridize for at least 30 minutes. 2 hours of pre-hybridization is preferred.

Hybridization

When there is 10-15 minutes left in the pre-hybridization, you should denature your DIG-labeled DNA probe (see DIG (non-radioactive) labeling of DNA if your probe is not yet labeled).
If this is the first time this probe is being used for hybridization, denature by boiling for 5 minutes and rapidly cooling in ice/water.
If the probe mix has been used before and is stored between ­150C and ­250C, then denature by heating in bath at 680C for 10 minutes before use.
When the pre-hybridization is complete, pour off the DIG Easy Hyb solution. Either pour in the sink or into a tube to be stored at 40C and used again.
Add 3.5 ml denatured DIG labeled DNA by pouring into the hybridization tube containing the filter.
Replace the tube in the Hybridization Incubator and start it running again (same temperature).
Filter should hybridize with probe for at least 4 hours and can go overnight.

Stringency Washes

83 Before you start the stringency washes start thawing (RT) a tube containing12.0 ml of 10X blocking solution. (This is started now because it takes ~ 40 minutes to thaw and is used in Immunological Detection after washes).
84 Make fresh wash solutions as follows:

2XSSC, 0.1% SDS (100 ml)

1. Add 10.0 ml 20XSSC stock to a clean, 100 ml graduated cylinder.
2. Add 1.0 ml 10% SDS stock to the graduated cylinder.
3. Add 89.0 ml dH2O to bring the liquid level to 100.0 ml.
4. Label the cylinder to avoid later confusion.

0.5XSSC, 0.1% SDS (100 ml)

1. Add 2.5 ml 20XSSC stock to a clean, 100 ml graduated cylinder.
2. Add 1.0 ml 10% SDS stock to the graduated cylinder.
3. Add 89.0 ml dH2O to bring the liquid level to 100.0 ml.
4. Label the cylinder to avoid later confusion.

Washes

1. Pour off the DIG labeled DNA probe from pre-hybridization into a plastic tube. Store the probe at ­150C to ­250C so that it can be reused.
2. WEARING GLOVES and using two pair of forceps, carefully remove the filter from the tube and place it in a small, plastic tub.
3. Rinse the hybridization tube out thoroughly with water.
4. Pour ~ 50.0 ml of 2XSSC, 0.1% SDS into the tray.
5. Place the tray on agitator at low-medium speed. Let the filter wash/agitate for 5 minutes at RT.
6. Meanwhile pour ~ 50.0 ml 0.5XSSC, 0.1% SDS into each of two hybridization tubes. Add the tubes to the hybridization incubator so that the tubes and solution will be pre-warmed to 550C.
7. After 5 minutes, pour off the first 50.0 ml of 2XSSC, 0.1% SDS. Add the remaining portion of 2XSSC, 0.1% SDS.
8. Replace the tray on the agitator for an additional 5 minutes of washing.
9. Turn off the agitator and pour off the solution.
10. Using forceps, replace the membrane properly (colony side up) in one of the hybridization tubes containing pre-heated 0.5XSSC, 0.1% SDS.
11. Wash filter at 550C for 15 minutes. Pour off.
12. Add the remaining 50.0 ml of 0.5XSSC, 0.1% SDS (from the balance tube) into the tube containing the filter. Add water to the second tube to balance this one.
13. Replace the tubes and wash at 550C for an additional 15 minutes.
14. Meanwhile make 100.0 ml of 1X Blocking Solution and 20.0 ml of Antibody Solution which you will need for Immunological Detection (directions below).
15. After 15 minutes, pour off the wash solution and proceed directly to Immunological Detection (below).

Making 1X Blocking Solution and Antibody Solution

You should have a tube with 12.0 ml of 10X Blocking solution which is thawed by now. If not, then thaw one.
Get 3 sterile, 50 ml tubes.
Add 45.0 ml maleic acid buffer to 2 of the tubes and label these 1X Blocking Solution
Add 18.0 ml maleic acid buffer to the last tube and label it Antibody Solution.
Make1X working solutions by adding 5.0 ml of thawed 10X blocking solution to the first 2 tubes and adding 2.0 ml of thawed 10X blocking solution to the last tube.
Mix thoroughly. The Blocking Solution is ready to use for Immunological Detection but the Antibody Solution needs further preparation:
Take out of the Lab Reagents 40C box.
Balance (400 ul balance tube) and centrifuge the anti-digoxigenin-AP (vial 4, stored at 4C) at 10,000 rpm for 5 minutes.
Add 2.0 ul of anti-digoxigenin-AP from the surface to the tube containing 20.0 ml blocking solution (making 1:10,000 dilution).
Mix thoroughly. The Antibody Solution is now ready to be used.

Immunological Detection

1. Leave the door of the hybridization incubator open and set the temperature to 240C to bring the incubator to RT. To help reduce the temperature rapidly a fan can be directed into the hyb/incubator. All the following steps should be performed at RT unless otherwise noted.
2. Add about 50.0 ml of Washing Buffer to the hybridization tube containing the filter.
3. Agitate and rinse briefly (5 minutes or less). Do this by hand or in the hybridization incubator if it has reached RT.
4. Pour off the Washing Buffer.
5. Add about 100.0 ml of Blocking Solution to the hybridization tube. Add 100.0 ml dH2O to the balance tube.
6. Incubate for 30 minutes at RT in hybridization incubator.
7. Pour off the Blocking Solution. Pour off the balance tube.
8. Add about 20.0 ml of Antibody Solution to the hybridization tube. Add 20.0 ml dH2O to the balance tube.
9. Incubate for 30 minutes at RT in hybridization incubator.
10. Pour off the Antibody Solution. Pour off the balance tube.
11. Add about 100.0 ml Washing Buffer to the hybridization tube. Add 100.0 ml ddH2O to the balance tube.
12. Incubate for 15 minutes at RT in hybridization incubator.
13. Pour off the Washing Buffer.
14. Again add about 100.0 ml Washing Buffer to the hybridization tube. The balance tube should have 100.0 ml ddH2O in it.
15. Incubate for 15 more minutes at RT in hybridization incubator.
16. Pour off the Washing Buffer. Pour off the balance tube.
17. Add about 20.0 ml Detection Buffer to the hybridization tube. Add 20.0 ml ddH2O to the balance tube.
18. Incubate for 2-5 minutes at RT in hybridization incubator.
19. Pour off the Detection Buffer and balance tube. Place the tubes in the tube rack.
20. Wearing gloves and using blunt forceps, carefully remove the filter and place it face up in a fresh development folder.
21. Lifting the folder so the top of the filter is exposed, apply 1.0 ml (4-5 drops) of CSPD ready-to-use (bottle 5, light sensitive, stored at 4C).
22. Immediately cover the filter with the top plastic sheet of the folder and spread the substrate evenly across the filter removing as many air-bubbles as possible.
23. Incubate at RT for 5 minutes.
24. Seal the development folder with tape to prevent drying of the filter (do this in such a manner that the tape does not block the exposure of any portion of the filter).
25. Incubate at 37C the filter for at least 10 minutes . (Increasing this step up to 1 hour is recommended as it will improve signal)
26. Expose the filter (face up) to X-ray film for 20 minutes. (See Exposing Film)
27. Develop the film (See Developing Film) and examine the results.
28. Test blots (smaller filters) can be stored as is (in their development folder) at 40C. Real library filters should be wrapped carefully in Saran Wrap, placed in a box for protection, and stored at -200C.

Stripping and Reprobing DNA Blots

1. If a filter is stored at 40C or -200C, then let it adjust to RT.
2. Remove the filter from its coverings.
3. Rinse the filter thoroughly in distilled water (a few minutes under a slow stream or shaking in a container filled with dH2O).
4. Using blunt forceps and wearing gloves, carefully place the blot face up in a hybridization tube.
5. Make 200.0 ml of 0.2M NaOH, 0.1% SDS in a 250 ml graduated cylinder as follows:
8.0 ml 5N NaOH
2.0 ml 10% SDS solution
190.0 ml ddH2O (bringing liquid to 200.0 ml total)
6. Add 100.0 ml 0.2M NaOH, 0.1% SDS to the hybridization tube containing the filter. Add an additional 100.0 ml 0.2M NaOH, 0.1% SDS to a balance hybridization tube.
7. Balance the tubes and rinse for 15 minutes at 37C in the hybridization incubator.
8. Pour off the solution from the tube containing the filter. Pour the 0.2M NaOH, 0.1% SDS from the balance tube into the tube with the filter.
9. Add 100.0 ml of dH2O to the balance tube.
10. Rinse for an additional 15 minutes at 37C in the hybridization incubator.
11. Make 100.0 ml of 2X SSC in a 100 ml graduated cylinder as follows:
10.0 ml of 20X SSC
90.0 ml of ddH2O
12. After the second rinse, pour off the solution from the tube containing the filter. To this tube add 100.0 ml of 2X SSC.
13. Balance tubes and rinse for 5 minutes at RT in the hybridization incubator.
14. Pour off SSC solution and water from balance tube.
15. At this point the filter is stripped and ready to be stored and/or reprobed. Store in 2XSSC or Maleic Acid Buffer at 40C. If you wish to reprobe, you can start immediately with pre-hybridization/hybridization.

DNA Quantification

Spectrophotometric Quantification of DNA

1. Turn on the spectrophotometer and allow it to heat up.
2. Prepare samples, including two Blanks, in sterile 1.5ml centrifuge tubes:
Blanks are 100 ul sterile ddH2O
Samples are 3ul DNA diluted in 97 ul
3. Clean (with dH2O) 2 quartz cuvettes (100 ul volume).
4. Pipet transfer Blank samples to the quartz cuvettes and "Auto Zero" the spec at the 260nm setting.
5. Clean the reading cuvette (front position)(with dH2O).
6. Add DNA sample.
7. Take reading and record the results.
8. Repeat the three previous steps for each remaining sample.
9. Clean the quartz cuvettes and put them away.
10. Turn off the spectrophotometer.
11. To calculate the concentration of your original samples, use the following formula:

Original DNA Sample Concentration (ng/ul) = (OD260/0.2) X (1000 ng/Dilution Factor)

0.2 is the standard value for 1 ug (1000 ng) of DNA
The Dilution Factor is the fraction by which you diluted your original sample

Acceptable concentrations of DNA samples vary based on the intended uses for the DNA. In general, though, a concentration of 100 ng/ul or higher is sufficient for most uses in lab.

For more information, see the Long Form.

Testing DNA Recovery using Agarose Plates

Making Gel Plates (only required if there are no plates available)

1. Make a 100 ml 1% agarose gel as normal (including 1XTAE Buffer, EtBr).
2. When the agarose is sufficiently cool pour it into 4 or 5 large, clean petri dishes.
3. Wait for these plates to solidify (about 30 minutes).
4. Label the plates (1% agarose gel) and use immediately or store at 40C.

Making DNA standards (only needed if standards are unavailable)

1. Bluescript is a good standard. Start from a stock, in this example PK SB+ (740ng/ul). From this stock, create serial dilutions in TE for final concentrations of 50, 25, 10, 5, 1, and 0 ng/ul.
2. Label 5, 0.6ml tubes appropriately.
3. The following chart shows dilutions for an original stock at 740ng/ul:

Final Concentration DNA amount/type added TE Volume Final Volume
50 ng/ul 13.5 ul stock (740ng/ul) 186.5ul 100ul
25 ng/ul 100 ul (50ng/ul) 100ul 100ul
10 ng/ul 100 ul (25ng/ul) 150ul 150ul
5 ng/ul 100 ul (10ng/ul) 100ul 150ul
1 ng/ul 50 ul (5ng/ul) 250ul 300ul
0 ng/ul NO DNA ADDED 100ul 100ul

Testing DNA Recovery

1. If necessary thaw DNA.
2. Make 3 dilutions of your DNA sample: 1/10, 1/100, 1/1,000. (2 ul sample in 18 ul sterile ddH2O)
3. WEARING GLOVES (because of EtBr), obtain a gel plate and use a straight edge and marker to construct a grid of boxes on the bottom of the plate (6 boxes for standard and 3 boxes for each sample).
4. Label boxes ("Standards"/sample name AND concentration/dilution factor).
5. Add 2 ul of each concentration of standard to its appropriately labeled box.
6. Add 2 ul of each concentration of sample to its appropriately labeled box.
7. Wait 5-10 minutes for the DNA to soak into the plate. Cracking the lid will speed up this step.
8. Photograph the plate as you would a gel (see Photographing a Gel).
9. Match your DNA sample dilutions to a standard dilution which is close in fluorescence.

(Example: I estimate that the 1/10 dilution of my DNA sample closely matches the 25 ng/ul standard. To calculate the concentration of my DNA prep I multiply:
(25 ng/ul) X ((1)/(1/10)) = 250 ng/ul)

For more information, see the Long Form.

Enzyme Reactions

Rules for using enzymes

1. Be careful with enzymes-keep them in a cooler outside of the freezer and use barrier tips!
2. If you use the last of an enzyme make sure to follow through on ordering more if it is needed. Always update the freezer list when new enzymes arrive.

Methods: Setting up enzyme reactions

1. Obtain wet ice and thaw template and other reagents if necessary.
2. Label properly one small tube for each reaction (96 well plates can also be used. In this case incubation is done in a 37 C incubator with parafilm covering the wells).
3. Turn on the 37 C water bath.
(PRE-MIX OPTION: If you are doing many reactions which are identical other than the template or enzyme you can make a pre-mix to save pipetting time. In a large tube add all common reagents. If you will use the pre-mix for 10 reactions, add 11 reactions worth of each reagent to allow for pipetting imprecision. Enzymes can be added to pre-mixes if they are shared in common with all of the reactions. In this case, add the enzyme last to the pre-mix)
4. Add the following reagents in the listed order (assumes a 10.0 ul reaction). (NOTE: Make sure to use buffers that match your enzymes. If an enzyme uses BSA in addition to its buffer, you will add 1.0 ul of 10X BSA and subtract 1.0 ul of ddH2O for a 10.0 ul reaction)
EXAMPLE REACTION:
7.75 ul sterile ddH2O
1.0 ul EcoRI Buffer
1.0 ul template (Amount may be varied depending on [ ] of template and desired application)
0.25 ul EcoRI (This is 5 units of actvitiy for this particular enzyme. 1 unit is able to cut 1ug of DNA in a 1 hour digest. Generally a 4-5 fold excess of enzyme is considered desirable)
5. When all reagents for the reaction have been added, mix the reactions by tapping them briefly. If needed, balance and spin down quickly.
6. Place the reaction tubes in a float in the 37 C water bath.
7. Let the enzyme digest for at least one hour for complete digest. Up to two hours or more can be better and should result in a complete digest under correct conditions. Set a timer to keep track if this is helpful.
8. After the digestion reaction is complete you may want to do one or more of the following (methods detailed below):
- Enzyme Inactivation
- Dephosphorylation
- Add 2.0 ul of 6X Loading Dye to a 10.0 ul reaction and run all 12.0 ul on a gel to confirm what bands are present or to cut out bands for further manipulation

For more information on general tips, selecting enzymes properly, buffer conditions, inactivation, dephosphorylation, and sequential or double digests, see the Long Form.

Making Freezing Stocks

1. Determine what you will freeze. For clones including unsequenced PCR fragments freeze 4 or more cultures to insure lower frequency of mutation.
2. First fill out the paperwork in the three notebooks and enter records into the computer database:

Fill out sample positions in "Freeze Box Maps" notebook

1. You will add your samples to the box in that series that is not yet filled (or start and number a new box if needed). Determine the appropriate series of box(es) for your samples. Here are the codes for the different types of boxes:
Box Letter Code Box Description
N Worm Strain in ­800 freezer
P Worm Strain in liquid nitrogen tank
Q Bacterial or Yeast Strain in ­800 freezer
R Primer in ­200 freezer
S Antibody in ­800 freezer
T DNA prep or other in ­200 freezer
U RNA prep
2. In the "Freeze Box Maps" notebook, find the sheet corresponding to the box you will use. Fill in sample information in the square that corresponds to where the sample will be stored in the box.

Fill out information in the "Strain" notebook

1. For bacterial and worm strain stocks you need to fill out the following information in the notebook: Strain Name, Genotype, Who prepared/froze it, Date frozen, and freeze position.

Fill out information in the "Plasmid/Cosmid Log" notebook
1. For plasmids and cosmids you will need to fill out the following information: Plasmid/Cosmid name, Selection Marker, CE, LG, or gene, Who prepped/froze it, Date frozen, and construction notes.

Enter stock information into the database

1. Open the desktop alias 'HMC database'.
2. File Maker Pro 4.1 will open, and a dialogue box will prompt for a password. Type "edit" and click "OK". (If you do not want to edit but only want to view records type "worm" for the password)
3. Press " N" for a new record (or select "New Record" under the "Mode" menu).
4. There are many possible areas to fill in for each new record. You will only need to fill in a few depending on your sample type.
FOR PLASMIDS/COSMIDS
1. Use the pull down bar to select "Bacterial Strain" for Item Type
2. Type the Strain name (as entered in the strain notebook)
3. Genotype (plasmid/cosmid name or code ­ as in strain notebook)
4. Selection marker (antibiotic used for selection)
5. LG, gene or protein (also as in the strain notebook)
6. AKA (additional ID information ­ prep number/letter, PCR number/identifier)
7. Host (bacterial host for transformation ­ XL1 Blue, for instance)
8. Description (notes on the plasmid/cosmid, including details of construction, PCR, etc.)
9. Source (who cloned the sample, where/who it was originally obtained from)
10. Date frozen (date the stock was frozen)
11. Address (Box/Position)
12. In edit mode, the records are automatically saved as changes are made. Continue to make new records and enter information until you are finished. Then you can exit the database by simply pressing " Q".
FOR WORM STRAINS
1. Use the pull down bar to select "Worm Strain" for Item Type.
2. Type the Strain Name.
3. Next to Genotype, type the genotype (mutations).
(NOTE: Information regarding the freezing of the strain is entered later by the technician handling the freeze/thaw process)
4. In edit mode, the records are automatically saved as changes are made. Continue to make new records and enter information until you are finished. Then you can exit the database by simply pressing " Q".

Making freezing stocks ­ Bacterial Strains

1. Obtain 1 clean, 1.8 ml freezer tube (colored tubes) for each sample.
2. Label the tubes with the following information ­ Strain name, Plasmid/Cosmid name, Selection Marker.
3. Make a 15% glycerol stock as follows:
4. Add 0.5 ml 30% glycerol to each freezer tube.
5. Add 0.5 ml of each liquid culture to its corresponding labeled freezer tube. Put the caps back on.
6. Take the freezing stocks to the ­800 freezer and put them in the correct box and position.
7. Add approximately 0.5 ml of bleach to each culture tube. Wait about 30 minutes for bacteria to die before you rinse down the drain. Rinse culture tubes thoroughly. Place them in the wire cage with other tubes that are drying. Once dry use 95% EtOH to remove any marker label still on the tube.

Making freezing stocks ­ Worm Strains

Pre-Freeze Steps

1. In order to freeze a worm strain you need to first grow either 2 large plates full of worms or 10-12 small plates. (If the strain is homozygous use large plates. If the strain is genetically complex (heterogenous, transgene animals) use small plates and pick individual animals.)
2. Let the worms grow on plates until they have just eaten all of the food (you don't want dauer stage or a plate where worms are burrowing. Having many worms at L1 stage is ideal). After you start a new plate it will take about 7 days (+ or ­ a 1-2 days) for the plate to get to this point. After 4-5 days you will want to monitor the status of the plates on a daily basis.
3. Place your worm plates in the "To be Frozen" box in the 150C incubator. Sign up your strain information on the freeze sheets and the lab technician will carry out the freezing protocol.

Freezing Protocol

1. Get wet ice.
2. Microwave the most current open bottle of freeze solution with agar for about 20 seconds. Leave the cap just barely on and watch closely for boil over. You do want the solution to boil for a few seconds in order to change all agar to liquid.
3. Let the freeze solution cool at RT until it is not molten but does not harden (the point at which it can safely be handled).
4. While the freeze solution is cooling, get a 15ml sterile polypropylene tube for each strain and label it with the strain name. Put these tubes on ice.
5. For each strain label 3 freezer tubes (label with strain name).
6. Light flame.
7. Get the most current, sterile bottle of M9 and flame the mouth.
One Strain at a time perform the following steps:
8. Pour about 2 to 3 ml of M9 into one of the worm plates. Tilt the plate to wash the worms down to one end.
9. Use a sterile pasteur pipet to repeatedly take up the liquid and reapply it to the plate in order to wash more worms down. Once most of the worms are washed (don't worry ­ you won't get them all), use the pipet to transfer worms + solution to the second plate.
10. Using pipet, repeat tilting and washing for the second plate.
11. Once the second plate (or in the case of small plates ­ the 12th plate) is washed thoroughly, use the pipet to transfer the worms + buffer to the appropriate 15 ml tube on ice. Transfer enough worm + buffer to bring the volume in the tube to 3.0 ml.
12. Repeat freeze steps (8-11) for each strain or until the freeze solution is cool enough to proceed with the next steps.
13. When the freeze solution is at a workable temperature (warm/not painful to touch the bottle) use a disposable 10ml plastic pipet to add an approximately equal volume (3.0 ml) of freeze solution to the equal volume (3.0 ml) of M9/worms in each 15 ml tube.
14. Cap tube and invert gently to mix the solution.
15. Use the same pipet from step 13 to add 1.0-1.5 ml of the mixture to each of the 3 appropriately labeled freeze tubes for that strain.
16. Throw out excess mixture and used pipets in a Sharps Box.
17. Repeat steps 13-16 for other strains. (NOTE: If the freeze solution starts to solidify you need to re-microwave it and wait for it to cool again.)
18. Place the freeze tubes in a styrofoam box at RT.
19. Take the freeze tubes to the ­700C freezer. In the top compartment you will find an area for storage of styrofoam boxes.
20. If you have not already done so, fill out the required information in the Freezing Notebook.

Test Thaw (3 days after freeze)
1. Label a plate with bacteria with the strain name.
2. Get ethanol burner, spatula, labeled plates, and freezer tube rack.
3. Take 1 of the 3 freezer tubes (­700C freezer) for the worm strain.
4. Light flame and sterilize a spatula.
5. Scoop out ~ 200-300 ul volume of the frozen strain and drop it on the plate, either on the bacteria or directly adjacent to it.
6. Sterilize the spatula.
7. Repeat steps 4-5 for any other test thaws.
8. Place plates (right side up so liquid does not spill) in the 200C incubator.
9. Throw out the freeze tubes (Sharps) with excess thawed worms.
10. Check the plates after 1 or more days to see how the worms survive the thaw. If there are many worms (100+) than the freeze and thaw can be judged successful. If the thaw was unsuccessful you will need to refreeze the strain. In either case, fill out a test thaw slip for the strains and return to the original submitter. They should check viable strains for proper genotype. Inviable strains should be resubmitted. Record test thaw dates in Freeze Notebook. Viable strains with confirmed genotypes can have permanent freeze positions assigned and information recorded in the database.

Thawing Frozen Stocks

Bacterial Strains

1. Obtain plates of the appropriate type (LB + antibiotic corresponding to the strain resistance).
2. Label bottom of each plate with bacterial strain name, your initials, and the date.
3. Use the database or strain notebook to locate the freeze position of the desired strain
4. Take the following to the ­700C freezer:
Ethanol burner
lighter
platinum inoculating loop
plates
styrofoam container
5. Light flame.
6. Find the freezing stock tube for you strain and place it in the styrofoam container (to slow any thawing).
7. Flame loop.
8. Touch the loop to the top of the frozen stock and move the loop in a smooth line across the surface of the first quarter of the plate.
9. Flame loop.
10. Streak again, starting from the end of the last streak and touching it only once. Streak out another quarter of the plate surface.
11. Repeat steps as above for the rest of the plate.
12. Replace the frozen stock in its proper position in the ­700C freezer.
13. Clean up.
14. Incubate plate at the proper temperature (usually 370C) for many hours. 10-16 hours is suggested for many bacteria.

For more, see the Long Form.

Worm Strains

1. Sign up on the list for thawing strains. The lab technician will thaw the strain and give you a plate in 1-2 days if the thaw is successful. It will then be your responsibility to maintain the strain.

Thawing Protocol

2. Label a worm plate with bacteria with the strain name and thaw date.
3. Use the database or strain notebook to locate the freeze position of the requested worm strain (the N is preferable as these strains are easily accessed from the ­700C freezer)
4. Take the following to the ­700C freezer:
Ethanol burner
Lighter
Ethanol bottle
spatula
labeled worm plates
styrofoam container
5. Light burner.
6. Place spatula in ethanol bottle.
7. Locate the frozen worm strain (N box) and place it in the styrofoam container to slow any thawing.
8. Flame spatula to sterilize.
9. Use the spatula to scoop out ~ 200-300 ul volume of the frozen strain and drop it on the plate, either on the bacteria or directly adjacent to it.
10. Sterilize the spatula.
11. Repeat steps if there are other strains to be thawed.
12. Replace the frozen strain(s) in their proper freeze positions in the ­700C freezer.
13. Clean up.
14. Check the thaw plate in 1 or more days to ensure that the strain has thawed viably. If the strain is viable, give the plate to the person who requested it. If the strain is not viable, you may need to thaw again or the frozen stock may be inviable.
(NOTE: If you thaw the last of a strain (i.e. no frozen stock remains), make absolutely sure to tell the recipient of the strain that they must resubmit the strain to be frozen again.)
(NOTE: Worm strains should have secondary tubes contained in liquid nitrogen (P Box). If the stock in the N Box runs out or is suspect, you can thaw from the stock in liquid nitrogen.)

Assigning Freeze Positions for Worm Strains

1. Freeze positions should be assigned to worm strains once test thaws have verified the viability and genotype of frozen stocks. By this point the basic information for the worm strain should be in the database.
2. Consult the freeze notebook to determine the position of the last worm strain to have a position assigned. For each strain, assign two tubes, one each to both a P Box position and an N Box position. Write the assigned positions for each strain into the following notebooks: "Freeze Box Positions", "Strain Notebook". Positions should also be added to the computer database records.
3. Move the worm tubes from where they are being stored in the ­700C to their assigned positions in the N Boxes and P boxes also found in the bottom of the ­700C.
(NOTE: If an N Box fills up then label and start a new one. These boxes are stored permanently at ­700C. If a P Box fills up then it is transferred to the liquid nitrogen tank (see below))

Transferring P Boxes to the liquid nitrogen tank

5. Take the full P Box from the ­700C freezer and bring into the liquid nitrogen tank.
6. Put on the blue, cold safety gloves.
7. Press the Reset Button on the tank lid (the green light should be ON before you remove the lid).
8. Remove the lid and set it aside.
9. Remove shelf number 4 or 6 (these are used for our lab storage) most of the way and hold it for a few seconds to let liquid nitrogen drain back into the tank.
10. Place the shelf on the floor.
11. Remove the guard.
12. Open the trap blocking the next available box position.
13. Place the P Box in the shelf.
14. Close the trap for that position.
15. Replace the guard.
16. Replace the whole shelf to its position in the liquid nitrogen tank (NOTE: this may take multiples attempts and an angled approach to succeed).
17. Press the Reset Button on the tank lid (the green light should be OFF before you replace the lid).

DNA Gel Electrophoreisis

Making 1X TAE from 50X stock (only needed when there is no 1X TAE left)

1. To dilute 50X TAE to 1X TAE do the following:
2. Measure 980 ml of dH2O in a 1000 ml graduated cylinder.
3. Open the 50X TAE (do not flame for sterility since TAE contains glacial acetic acid) and draw or pour 20 ml into the graduated cylinder to bring the liquid level to 1000 ml.
4. Pour this mixture into a clean, 1.0 L screw top bottle.
5. Screw the tops on the bottles. Mix the 1X TAE.
6. Label it as follows:
1X TAE (the date it was made) (your name or initials)

Making loading dye/ladder solution from stock
(NOTE: ladder/dye is normally stored at 40C in Lab Reagents Box, if not there are usually new tubes in the ­200C Lab Reagents Box)

NEB Ladder

1. For each tube make 1.0 ml of solution that is 1X Loading Buffer and 20ng/ul of DNA Ladder. Use the following amounts and reagents:

793.4 ul sterile (autoclaved) water
166.6 ul 6X Loading Dye
40.0 ul 500 ng/ul NEB DNA Ladder
1.0 ml loading dye/ladder solution
2. Get wet ice and thaw the DNA Ladder on ice (about 30 minute wait time).
3. Label a sterile (autoclaved) 1.5 ml tube as NEB Loading dye/Ladder solution and add the date and your initials.
4. Add the reagents in the amounts and order shown above. Mix, spin down and place in the Lab Reagents Box at ­200C (or if using immediately store at 40C).

Fermentas (1 kb DNA Ladder)

Get wet ice and thaw the DNA Ladder on ice (about 20 minute wait time).
Label 5, sterile (autoclaved) 1.5 ml tubes as Fermentas Loading dye/Ladder solution and add the date and your initials.
To each of the 5 tubes add the following reagents in order and amount:

396.7 ul sterile (autoclaved) ddH2O
84.3 ul 6X Loading Dye
20.0 ul 500 ng/ul Fermentas DNA Ladder

Mix, spin down and place in the Lab Reagents Box at ­200C (or if using immediately store at 40C).

Making agarose gel (50ml, 1%)

There are different grades of agarose that can be used in gels and gels can be made as varying percentages in solution.

Gels are made by the same basic procedure despite differences in volume of gel and amount, type and percentage of agarose:

1. Put on gloves (to avoid EtBR contamination).
2. Place a clean, 250 ml flask on the benchtop.
3. Weigh out 0.50 g of Low EEO agarose and carefully add it to the flask.
4. In a graduated cylinder measure 50 ml of 1X TAE solution. Pour this solution into the flask containing the agarose and swirl gently to mix.
5. Place the flask in the microwave with a paper towel over the top.
6. Microwave the gel for 1 minute or until agarose is completely dissolved.
7. Let the gel cool until you are able to touch the flask without pain (5-15 minutes).
8. Wearing gloves and using a P-20 add 2.5 ul of 10mg/ml EtBr. NOTE: EtBr is a strong mutagen (carcinogenic) so make sure to wear gloves and handle properly. Tips go in the sharps box! From this point on, wear gloves when handling the gel or apparatus.
9. Swirl the flask gently (you do not want too many air bubbles) to distribute the EtBr.
10. Set up the tray for the pour. Make sure things are straight, even and tight. You may either leave the comb in or take it out and add it later.
11. Pour the gel. Do not forget to add the comb if you left it out initially!
12. Wait 15-20 minutes for the gel to set up.
13. Clean out the 250 ml flask (rinse it) and return it to the area since it will be used to make future gels.
14. When the gel is set you can remove the comb. It is helpful to add a little1X TAE buffer around the comb to facilitate its removal. Carefully work the comb out (use side to side rocking while gently pulling up), trying not to damage any of the wells.
15. Remove the tray with the gel and place it in the gel apparatus. The end of the gel with wells should be toward the black terminal of the apparatus(a hole in the left side of the gel holder will slide into place in the apparatus).
16. Pour 1X TAE buffer into the apparatus until it just covers the gel and fills the wells.

For more on agarose types and varying gel compositions, see the Long Form.

Making samples

-From Enzyme digest reactions
5. It is common to run your complete digest on a gel (unless you will be using liquid digest for further applications).
6. Add 6X Loading dye to each reaction in a volume that will make a 1X sample (Ex: Add 2 ul dye to a 10 ul reaction, 4 ul dye to a 20 ul reaction).
7. Once the samples are prepared and the gel is in place in the buffer (see above) then you can load the wells of the gel.

-From PCR Reactions

1. Write labels for your reactions on either small tubes, a 96 well plate, or a strip of parafilm. Add 10% of your reaction volume (5.0 ul for a 50 ul reaction) to the appropriately labeled tube (NOTE: For 25 ul reactions it is recommended to take 4 ul for your sample). Add 1.0 ul 6X Loading Dye (to make 1X dye) to each of these samples.
2. Once the samples are prepared and the gel is in place in the buffer (see above) then you can load the wells of the gel.

Loading the gel

1. Load from left to right and keep careful track (in your notebook) of the samples in each well. It is customary to load 10ul of dye/DNA ladder in the leftmost well (this will be used as a standard for comparison of size and intensity).

For more advice on loading the gel, see the Long Form.

Running the gel electrophoreisis apparatus

1. Once the gel is immersed in buffer, and loaded and positioned properly (wells towards the black terminal), place the lid securely over top [make sure that black matches black (upper right corner)].
2. Make sure the power source is off (grey switch flipped to O).
3. Connect the leads to the power source (match red with red and black with black).
4. Flip the grey switch on the power source to turn it on (flip to I).
5. Flip the Milliamps/Volts switch to check for current (milliamps) or voltage (volts).
6. Run most gels between 80 and 100 volts (do not exceed 100 volts!).
(NOTE: Roughly speaking, voltage is proportional to speed of gel migration and inversely proportional to resolution of bands)
When the gel is running you should observe bubbles forming around the encased platinum wire near the black terminal. With time gel bands should migrate forward (toward red) from the wells.
7. Before you remove your gloves clean up the gel and sample materials.
8. Keep track of the gel by checking it occasionally (you do not want your bands to run off the gel). Gels run sufficiently in 1 hour or less at most voltages.

Photographing a gel

1. When your gel has run long enough turn the power supply off and disconnect the leads.
(There are two imaging areas available for photographing gels: Room 225 and Room 915)

Room 225 ­ Johnson Lab Imaging Equipment

2. Because of regulations we are unable to wear gloves in the elevator. I recommend putting your hands part way into gloves in order to safely handle and carry EtBr (only use non-gloved fingers to press elevator buttons). I also recommend wearing a lab coat.
3. Remove the gel and its tray and place both inside a "Gels Only" container.
4. Take gel container, gloves, lab coat and face shield to the elevator by Helen's office. Take the elevator to the 2nd floor. Go down the hallway about halfway to room 225 (Johnson Lab). There is a small alcove in this entry room to 225 where gels are photographed.
5. Put on gloves fully.
6. Check the following items:
7. Monitor power is ON.
8. Printer power is ON.
9. Printer switch position is "down".
10. Lens cap is OFF.
11. Place your gel on the UV light box.
12. In order to position the gel as you want it, it is most convenient to have the camera set for real time. If it is not already set this way, you can reach this status by clicking on the camera icon.
13. Change the lens height and position if desired (with the hand cranks).
14. Focus the camera on your wells (adjusted by turning focus at the bottom of the lens).
15. Put on the visor/face shield to protect you from UV light.
16. Pull the door closed to limit the amount of white light.
17. Turn ON the UV light box.
18. Change the exposure time as desired to obtain a good picture(done by clicking on the arrows in the dialog box). Keep in mind the strengths of the bands relative to each other and the comparisons you want to make.
19. Once you are satisfied with the image click on "Done".
20. Turn OFF the UV light box.
21. Print your picture (click on the printer icon) and tear it off.
22. When satisfied with picture(s) do the following: turn off monitor and printer power, flip the printer switch up to the middle position, return your gel to its container, wipe off the light box, throw our your gloves, return to lab w/picture and gel (you may want to use lab coat as a barrier between you and EtBr contaminated materials).
23. See below for Interpretation of Gel Results after returning to lab.

Room 915 ­ Chang Lab Imaging Equipment

5. Put on gloves.
6. Remove the gel and its tray and place both inside a "Gels Only" container.
7. Go to 915.
8. Place gel on UV box and close the lid.
9. Check to see that the monitor power is ON and the lens cap is OFF.
10. Turn off lights as you feel necessary. Press "Record Image" to capture an image. Exposure time is adjusted with arrows on the control apparatus ("Darker" and "Brighter"). Focus is adjusted by turning at the base of the camera. Zoom can be adjusted on the camera (turn via a protruding knob).
11. To print the displayed image, press "Print Image" and tear off your printed picture.
12. When satisfied with your picture(s), do the following: clean off the gel box, place lens cap back ON, turn the monitor power OFF, return to lab with your gel and picture.
13. See below for Interpretation of Gel Results after returning to lab.

Interpreting Gel Results

1. Interpretation of your gel depends on the nature of the samples you are running. With different samples you will be looking for different size bands with different intensities so determination of expected bands ahead of time is crucial to successful interpretation. In general, here a few things to consider:
Make sure all lanes are properly labeled on your picture or in your notebook (name of sample, amount loaded, etc.).
Find your ladder and locate crucial bands for size comparisons. Use these bands (along with a straight edge if this is helpful) to estimate the size of your sample band(s).
Compare the brightness of bands with each other and with ladder bands. You can estimate the concentration of bands based on the following: 1. 10 ul of 20ng/ul ladder = 200 ng, 2. Intensity for a given [ ] of DNA varies with size (Thus a smaller band of the same [ ] looks fainter than a larger band of the same [ ])
Are there single, clear bands standing alone or is there background or other fainter bands? Single clear bands are expected from good PCR reactions. Many bands indicates a background of many products or digest fragments. Doublets occur when multiple bands of a similar size are present --this causes a thicker, brighter band.

After interpretation there are two situations:
18. You have the information you need from your gel. In this case, dispose of your gel in a burn box. Rinse the gel tray and container with tap water and place them back in the gel area. Wash your hands and tape your gel picture(s) into your notebook and label it.
19. You are unsure of your results or have not finished with the gel. In this case, there are many options:
Your experiment did not work and you need to repeat it with different conditions.
Run the gel further to better resolve bands.
Load more sample volume on a new gel.
Cut specific bands for other applications (ligation/cloning, reamplification)
Get a second opinion from someone else on interpretation of the gel
Wrap the gel in Saran Wrap, label it with date, initials and buffer, and store it at 40C until you may reuse it (NOTE: bands will diffuse over time).

Gel Band Cutting

Theory: Running samples and digests on a gel and then cutting specific bands allows you to isolate inserts, vectors, and other pieces of DNA. We use normal gels and a long wavelength UV handheld lamp to observe and cut the bands. Other techniques may employ low melt gels and light boxes or lamps of varying wavelength.

1. Make a normal DNA gel with enough wells to run your samples plus one or two control ladders to help you estimate band size. To prevent cross contamination between samples you may leave empty wells between them when you load the gel.
2. Load your samples and run the gel (running the gel at a lower voltage may help give the bands better definition). NOTE: Load 20 ul of DNA Ladder so that it will be clearly visible under the UV handheld lamp.
3. It is critically important to figure out what bands will be present when you run your samples on a gel. This will ensure that you know which bands to cut and which to leave behind.
4. Gather the following equipment which you will need to cut bands:
a piece of Saran Wrap (about 12"X12" or larger)
a labeled, sterile 1.5 ml tube for each gel band to be cut
a rack to hold your tube(s)
a spatula (the smaller version works best, found in drawer under balance) to scoop up bands
a razor blade to cut the bands (found in drawer under balance)
Face shield
Lab Coat (if desired for protection from UV)
Handheld UV lamp
5. When you feel the gel has run for a sufficient time to isolate bands, turn off the gel apparatus.
6. Place the gel without tray on the sheet of Saran Wrap.
7. Take the gel and equipment to a dark area where the bands can be visualized and cut (Room 932 is usually good).
8. Arrange materials, open the tube(s) that will hold the cut bands, put on your face shield, and plug in the UV lamp. To turn on the UV lamp press the black button.
9. Find the band you want to cut. Carefully use the razor blade to incise the gel around the band on all 4 sides. (If you do not see your band you can "cut blindly" in the expected region and hope that you got some product to use or to reamplify) (Try to get all of your band but not any part of another band. Surrounding (empty) gel is allowable but may be trimmed off)
10. Use the end of the spatula to bring the cut band out of the gel. If desired, put the band down on the plastic wrap and trim away any excess (non-band) gel.
11. Use the spatula to drop the band into the properly labeled tube. Close the tube.
12. Repeat Steps 9-11 for all the bands you want to cut.
13. Return materials to their proper areas. Dispose of the following: razor blade into sharps box, gel and plastic wrap into biohazardous waste box (save the gel if you want to check your gel after cutting to see what you missed or where you cut). Wash off the spatula and set it to dry.
14. Cut bands in their tubes can be stored at 40C or you can proceed directly to Gel Purification. To purify the cut bands for other uses follow the instructions for the protocol below (QIAQuick Gel Extraction Kit Protocol).

QIAQuick Gel Extraction Kit Protocol
(adapted from Qiagen protocol book page 24)

1. This protocol is adapted from the QIAquick Spin Handbook, page 24 and you may want to use that as a reference when performing this protocol.
2. Open the QIAquick PCR Purification Kit (Catalog No. 28704).
3. Check to see that a bottle of Buffer PE has ethanol added by looking to see that it is checked on the lid.
4. Turn the heating block on and set it to heat to 500C.
5. Weigh and record your gel cuts. Do this by taring the balance with an empty tube and then placing your tube with gel cut on the balance.
6. Add 3 gel volumes of Buffer QG to each gel cut.
(For this calculation assume that 100mg 100ul. Example: If your gel cut weighs 0.150 g, then you would add 450 ul of Buffer QG)
7. Incubate the gel tubes at 500C (block) for 10 minutes to completely dissolve the gel. Vortex each tube every 2-3 minutes to aid in dissolution.
8. Check to see if the liquid is yellow in color. [NOTE: Sometimes bands are blue from loading dye ­ this is fine] [If liquid is orange or violet the pH has changed too much. In this case, add 10 ul of 3M KAc (pH 5) and mix. The solution should then turn yellow]
9. Add 1 gel volume of isopropanol (Example: Gel cut weighs 0.150g, then add 150 ul of isopropanol)
10. Place spin columns from Qiagen Kit (labeled for each of your samples) into 2 ml collection tubes.
11. Using a pipetter to transfer your gel sample to the appropriately labeled spin column. (NOTE: The columns hold a maximum of 800 ul. If this is insufficient then add only 800 ul, spin, then add the rest to the column and spin again)
12. Balance tubes and spin for 1 minute at maximum speed.
13. Discard the flow through into a sink of running water (Do not rinse tubes at all!)
14. [Preferred Optional Step] Add 0.5 ml Buffer QG to each column and centrifuge at maximum speed for 1 minute. (This step removes traces of agarose and should be done when DNA will be subsequently used for direct sequencing, in vitro transcription, or microinjection)
15. To wash, add 0.75 ml of Buffer PE to columns. (NOTE: If DNA is being used for salt sensitive applications, like blunt-ended ligation and direct sequencing, let it stand for 5 minutes before continuing)
16. Balance tubes and centrifuge at maximum speed for 1 minute.
17. Discard the flow through into a sink of running water (Do not rinse tubes at all!)
18. Balance tubes and centrifuge at maximum speed for 1 additional minute. (to remove residual EtOH)
19. Discard the flow through into a sink of running water (Do not rinse tubes at all!)
20. Throw out collection tubes in burn box or sharps box.
21. Add spin columns to sterile, labeled 1.5 ml tubes.
22. Add 30 ul or 50 ul of Buffer EB (10mM Tris, pH 8.5) to the center of the membrane of each spin column. (The amount added depends on the desired final volume and concentration of your samples)
23. Let the columns stand for 1 full minute to increase recovery.
24. Balance tubes and centrifuge 1 minute at maximum speed to elute DNA.
(The final volume will be about 28 ul or 48 ul depending on which volume of Buffer you used to elute your sample. Recovery over Qiagen columns is 70-80% of original DNA with a maximum column binding capacity of 10 ug. Fragment size may range from 70bp ­ 10 kb)
25. Purified products may be used for further applications (ligation, sequencing, etc.) You may want to take a spec reading to verify sample concentration or run a gel to test the recovery of your samples. Sample make up (for a 28ul final volume) could be as follows:

Samples Vector (if included)
5 ul gel purified DNA 1 ul Vector

QIAQuick Gel Extraction Kit Protocol
(adapted from Qiagen protocol book page 24)

1. This protocol is adapted from the QIAquick Spin Handbook, page 24 and you may want to use that as a reference when performing this protocol.
2. Open the QIAquick PCR Purification Kit (Catalog No. 28704).
3. Check to see that a bottle of Buffer PE has ethanol added by looking to see that it is checked on the lid.
4. Turn the heating block on and set it to heat to 500C.
5. Weigh and record your gel cuts. Do this by taring the balance with an empty tube and then placing your tube with gel cut on the balance.
6. Add 3 gel volumes of Buffer QG to each gel cut.
(For this calculation assume that 100mg 100ul. Example: If your gel cut weighs 0.150 g, then you would add 450 ul of Buffer QG)
7. Incubate the gel tubes at 500C (block) for 10 minutes to completely dissolve the gel. Vortex each tube every 2-3 minutes to aid in dissolution.
8. Check to see if the liquid is yellow in color. [NOTE: Sometimes bands are blue from loading dye ­ this is fine] [If liquid is orange or violet the pH has changed too much. In this case, add 10 ul of 3M KAc (pH 5) and mix. The solution should then turn yellow]
9. Add 1 gel volume of isopropanol (Example: Gel cut weighs 0.150g, then add 150 ul of isopropanol)
10. Place spin columns from Qiagen Kit (labeled for each of your samples) into 2 ml collection tubes.
11. Using a pipetter to transfer your gel sample to the appropriately labeled spin column. (NOTE: The columns hold a maximum of 800 ul. If this is insufficient then add only 800 ul, spin, then add the rest to the column and spin again)
12. Balance tubes and spin for 1 minute at maximum speed.
13. Discard the flow through into a sink of running water (Do not rinse tubes at all!)
14. [Preferred Optional Step] Add 0.5 ml Buffer QG to each column and centrifuge at maximum speed for 1 minute. (This step removes traces of agarose and should be done when DNA will be subsequently used for direct sequencing, in vitro transcription, or microinjection)
15. To wash, add 0.75 ml of Buffer PE to columns. (NOTE: If DNA is being used for salt sensitive applications, like blunt-ended ligation and direct sequencing, let it stand for 5 minutes before continuing)
16. Balance tubes and centrifuge at maximum speed for 1 minute.
17. Discard the flow through into a sink of running water (Do not rinse tubes at all!)
18. Balance tubes and centrifuge at maximum speed for 1 additional minute. (to remove residual EtOH)
19. Discard the flow through into a sink of running water (Do not rinse tubes at all!)
20. Throw out collection tubes in burn box or sharps box.
21. Add spin columns to sterile, labeled 1.5 ml tubes.
22. Add 30 ul or 50 ul of Buffer EB (10mM Tris, pH 8.5) to the center of the membrane of each spin column. (The amount added depends on the desired final volume and concentration of your samples)
23. Let the columns stand for 1 full minute to increase recovery.
24. Balance tubes and centrifuge 1 minute at maximum speed to elute DNA.
(The final volume will be about 28 ul or 48 ul depending on which volume of Buffer you used to elute your sample. Recovery over Qiagen columns is 70-80% of original DNA with a maximum column binding capacity of 10 ug. Fragment size may range from 70bp ­ 10 kb)
25. Purified products may be used for further applications (ligation, sequencing, etc.) You may want to take a spec reading to verify sample concentration or run a gel to test the recovery of your samples. Sample make up (for a 28ul final volume) could be as follows:

Samples Vector (if included)
5 ul gel purified DNA 1 ul Vector
1 ul 6X loading dye 1 ul 6X loading dye
4 ul TE

Developing Film

1. The developer needs to warm up about 5 minutes before it is ready to develop film. If you have not already turned the developer (Room 915) do so.
2. Once the developer has warmed up, take the autoradiography cassette (with exposed film) to the Dark Room (Room 915).
3. Turn off any lights (you may leave the red light on if desired).
4. Turn off the room light and lock the door.
5. Open the cassette and take out the film.
6. Open the lid to the developer and slide the film (face up) in the hole at the back. You should feel the developer take the film and pull it in.
7. Wait exactly 2 minutes for the film to be developed. It will spit out into the bin on the floor.
8. Turn on the room light and unlock the door.
9. Look at the film to observe the results.
10. Turn off the developer if you are done with it.

Exposing Film

1. Take the appropriately sized Autoradiography Cassette, the box of individually wrapped Imaging Film (X-OMAT AR), and whatever you are exposing to the Dark Room (Room 915).
2. Turn off any lights (you may leave red light on if desired).
3. Turn off the room light and lock the door.
4. Open the autoradiography cassette.
5. Place the development folder (containing what you want to expose) in the autoradiography cassette with signal side facing up.
6. Open the film box and take out a single, individually wrapped sheet of film. Make sure you get only one piece of film.
7. Tear open the wrapper by pulling down the right side.
8. Take the film out of the paper sheath that contains the it.
9. Place the film over the development folder (in the autoradiography cassette). If desired crease one corner of film for help in orientation later.
10. Carefully close the cassette so that it seals completely over the film. Check to make sure the felt around the edges completely blocks light.
11. Seal the cassette by properly closing the fasteners.
12. The film is now being exposed. Start a timer to keep track of how long the film is exposed.
13. Check to make sure the box of individually wrapped film is closed!
14. Open the door and return to lab to wait during exposure time.
15. About five minutes before the film will be done exposure, return to the dark room and turn on the developer.
16. Once the film is finished being exposed, develop it (See Developing Film)

Inoculating LB +Op

Background: LB + Op bacteria is a lab strain of E. coli. We apply it to worm nutrient plates to provide food for worms. To obtain this media bottle of LB are inoculated with the bacterial strain.

There are two methods for inoculating:
1. Use liquid from the bottom of the old bottom to inoculate the new bottle;
2. Grow and/or pick an LB +Op colony and use it to inoculate the new bottle.

The first method is used most. The second method should be done occasionally to ensure purity or when you fear there has been contamination.

With either method sterile technique is very important because this is a point where you can introduce outside contaminants that may ruin many plates and experiments.

Method 1: Using liquid from the old bottle to inoculate the new bottle
1. Obtain an unopened, sterile 250ml bottle of LB. To the label, add "+Op" and the current date.
2. Stick green plunger or Pipetman on the end of a sterile, 10ml glass pipet (but do not remove it from the sterile metal sleeve yet).
3. Light flame
Open the new bottle of LB and flame the mouth and cap. Open the old bottle of LB +Op and flame the mouth and cap. Remove the pipet from the sterile metal sleeve and flame it.
Using the pipet, take just a drop or two from the old bottle. Add this to the new bottle.
Flame both bottles and caps. Screw the caps back on.
Turn the flame off now.
Let the new bottle of LB +Op sit at RT for at least 24 hours to allow the bacteria to grow.
Clean the old bottle as follows: add some 10% bleach solution, wait a few minutes, rinse thoroughly down the drain, wash bottle with soap and water, set to dry.
After a day, the LB +Op should be ready to test on about 20 plates (See Spreading Bacteria on Plates for method). Wait a day for the bacteria to grow and observe the plates to see if any other bacteria seem to have grown. If the plates look normal then you can assume the bottle is good and use it freely. If the plates show signs of contamination you will have to re-inoculate using the second method.
LB + Op is stored at 40C when it is not being used.

Method 2: Growing and/or picking an LB +Op colony to inoculate the new bottle
1. Streak and grow a plate of LB +Op from stock unless there is a recent plate stored at 40C (see Streaking and Incubating Plates).
2. Get an unopened, sterile 250ml bottle of LB. To the label, add "+Op" and date.
3. Get an inoculating loop and your LB +Op plate.
4. Light flame.
5. Open the new bottle of LB and flame the mouth and cap.
6. Flame the inoculating loop and use it to pick one colony from your plate.
7. Dip the loop in the bottle of LB briefly.
8. Remove the inoculating loop and flame the mouth and cap of the bottle.
9. Screw the cap on.
10. Flame the inoculating loop to sterilize it.
11. Let the new bottle of LB +Op sit to grow. At RT it will take 2 days to grow (NOTE: This is longer than the first method). In the 37o incubator the LB +Op will grow in 1 day.
12. When it is grown the LB +Op is ready to test on about 20 plates (See Spreading Bacteria on Plates for method). Wait a day for the bacteria to grow and observe the plates to see if any other bacteria seem to have grown. If the plates look normal then you can assume the bottle is good and use it freely. If the plates show signs of contamination you need to re-inoculate.
13. LB + Op is stored at 40C when it is not being used.

Ligation Reactions

Theory: DNA strands with like ends (either sticky or blunt) can be brought together in a reaction using T4 Ligase enzyme. Templates should be carefully designed and digested to avoid self-ligation (using phoshpatase, especially on vectors, helps with this) or ligation in multiple orientations leading to unwanted clones. It is ideal that templates are double cut with sticky but non-compatible enzymes.

1. First, design your ligation reactions and write out the volumes of reagents you will use.

[NOTE: Most ligation reactions work best when there is approximately a 3:1 ratio of vector:insert, taking into account relative concentrations and molecule sizes. Start by choosing a vector concentration in the reaction (usually about 1000ng). The calculation is demonstrated in the following formula and example:

Amount of insert (ng) = Amount of vector (ng) X 3 (desired ratio)
Ratio of vector band size:insert band size

The following example is for a 4.5 kb vector to be added at 1500 ng in a ligation reaction with a 0.8 kb insert.

Amount of insert (ng) = 1500 ng X 3 = 4500 ng = 750 ng
4.5:0.8 6

Thus, if the vector is 750 ng/ul in concentration and the insert is 75 ng/ul in concentration, you will add 2 ul vector and 10 ul insert to ligation reactions.

2. For each type of ligation, you should also run two controls. The first control should be insert alone. The second control should be vector alone. (NOTE: If you are running multiple inserts with a similar vector, you need only include one control for the vector) The following recipes are standard though levels of template can be varied:

Insert Alone Vector Alone Vector + Insert
7.75 ul ddH2O 16.75 ul ddH2O 6.75 ul ddH2O
2.0 ul Ligation Buffer 2.0 ul Ligation Buffer 2.0 ul Ligation Buffer
10.0 ul Insert 1.0 ul Vector 10.0 ul Insert
0.25 ul T4 Ligase 0.25 ul T4 Ligase 1.0 ul Vector
0.25 ul T4 Ligase

3. Label a 0.6 ml, sterile tube for each reaction. Place the tubes in a rack. Add the reagents in order (from top to bottom) as shown above. Add the Ligase (found in the enzyme freezer) last and take an aliquot using a barrier tip. Draw from this aliquot to add to your reactions.
4. When all reagents are added tap the tube to mix.
5. Place the tubes (in the rack) in the 150 incubator.
6. Let the ligation reactions run for a minimum of 1-2 hours. The efficiency of the reaction increases almost linearly with time, so it is ideal to let the ligations run from 4 hours to O/N before transforming.
7. Follow the protocol for Transformation to determine the results of the ligation reactions.

Making Small Liquid Cultures (5.0 ml)

Making LB with antibiotic (you may need to do this)

1. Thaw the appropriate antibiotic (depending on the one used in your plates) on ice, picking a tube already opened if there is one. If you only use part of the antibiotic in a new tube you should label it to indicate that is has been opened. Antibiotics are stored at -200C in a box labeled "Antibiotics".
2. Obtain a sterile bottle of LB and estimate the volume of LB. (Ex. ~135 ml)
3. Add antibiotic to the LB. The quantity you add will depend on the concentration of the antibiotic and the volume of the LB. You want a final concentration of 1ug/1ml of antibiotic in LB. Example:

50mg/ml antibiotic stock: Add 135ul carb to 135ml LB

4. Swirl the LB bottle to mix.
5. Label it with the antibiotic, its color code, and the date.
6. When it is not being used it should be stored at 40C.

Making Liquid Cultures
It is essential to match your liquid antibiotic with the antibiotic of the plate you pick from. Once you have an adequate volume of LB w/antibiotic to make your cultures you can proceed:
1. Label one sterile glass tube for each prep. If you are picking multiple colonies from a single plate indicate this by adding a series of letters to your labels. (For example, a series of tubes for cultures from a plate called 3.1c might be labeled as follows: 3.1cA, 3.1cB, 3.1cC, 3.1cD, etc.)
2. Sterile technique is important to prevent contaminating your culture. Fill a plastic or glass pipet with LB + antibiotic and dispense 5 ml to each of your culture tubes.
3. Get forceps and sterile (autoclaved) toothpicks. Get the plates containing the colonies that you want to prep.
4. There are two methods for picking a colony and adding it to the LB/antibiotic:

Preferred Method. Light the burner. Flame the tips of the forceps. Remove the cap from the toothpick tube. Flame the mouth of the toothpick tube. Use the forceps to pick out one of the toothpicks. Flame the mouth of toothpick tube. Place the cap back on the toothpick tube. Touch the toothpick (using the forceps to manipulate it) to a single colony on your plate in order to pick it. Open the cap to the appropriate culture tube. Flame the mouth of the culture tube. Drop the toothpick in. Flame the mouth of the culture tube. Place the cap back on the culture tube (fairly tight although not all the way down).

For Method 2 (where toothpicks are not added to culture tubes) see the Long Form.

Regardless of which method you use, make sure to handle toothpicks in a sterile manner. If you drop or contaminate one simply throw it out and try again with a sterile toothpick.
5. When all the liquid cultures are inoculated, take the culture tubes to the Incubator Shaker (Room 968) and place them for overnight growth (or 16 hours).
6. Cut strips of parafilm and stretch them around your plates to seal them. Store the plates at 40C.
7. Retrieve the cultures from the Incubator Shaker the next day (or after 16 hours). Use them for preps of other applications and store them at 40C when they are not being used.

PCR (Polymerase Chain Reaction)

Theory: PCR is a cycled reaction that allows for ultimate amplification of DNA products. Taq or other polymerases carry out the synthesis portion of the reaction. The primary stages of the reaction are: 1. DNA is melted (940C) to single-stranded state, 2. Primer is annealed (reaction specific temp) to DNA, 3. Synthesis is carried out (usually 720C). The cycle of 3 steps is repeated 25-35 times to result in exponential amplification.

Preparing for PCR

Ensure that you have the reagents you need for the reaction ­ proper and sufficient template, nucleotides, primers, buffer, polymerase, etc.
Sign up on the PCR sign up sheet.

Making Nucleotide Stock

NOTE: These steps are not performed every time. You need only do this when you have run out of 10X dNTPs or feel that the old nucleotides are degraded.

1. Thaw the set of dNTPs on ice (they are found in a small box in the enzyme freezer). This will take about 20 minutes.
[You want to make a dNTPs stock where each nucleotide is 2mM. Since the stocks dATP, dCTP, dGTP, dTTP are 0.2 mM, you will make each10X in solution. The total end solution will be 8mM (4 X 2mM)]
2. To a sterile 1.5 ml tube add 184 ul of TE.
3. To this, in turn, using barrier tips, add 4 ul of dATP, dCTP, dGTP, and dTTP.
4. This makes a final volume of 200 ul. Label the tube 10X dNTPs (2mM each) and store it at -200C when it is not being used.

Tips on Primer Design

In general:
Higher G/C content makes for more stable and effective primers because the G/C bond is higher in energy than the A/T one
To estimate annealing temperature for a primer, use web site calculators or the following estimate: 40C for each G or C and 20C for each A or T (when designing multiple primers for identical reactions you may want to keep their estimated annealing temperatures to a close range)
Primers should be 20 bases or greater in length (more length equals more stability and specificity)
The 3' end of the primer is crucial since this will bind first
Avoid excessive self-dimers and hairpins, especially those that form high-energy bounds. These primer-primer interactions can tie up primer and prevent it from binding template properly. To check for these interactions use on-line diagnostic programs.
Enzyme sites for cloning and diagnostics should be added to the 5' end of the primer. In this case it is useful to include an additional three bases in the primer, 5' to the enzyme sequence, in order to stabilize the primer. These additional bases do not need to match template; choose whatever fits best.
If you can not find a potential primer in a given sequence area of your template, try looking in a different area. If a particular area is causing trouble (hairpin/self-dimer) try shifting the primer sequence to avoid or partially cut off that area.

Degenerate Primers:

Degenerate primers contain a mixture of possible primer variations. These oligos allow for variation at specific bases where template sequence is unknown. The most important consideration in designing degenerate primers is the degree of degeneracy of the primer.
The following are the IUB codes for mixed base sites:

N = G, A, T, C V = G, A, C B = G, T, C H = A, T, C D = G, A,T
S = G, C W = A, T K = G, T M = A, C Y = C, T
R = A, G

An example of a degenerate primer sequence follows:
TAYGGN (where four variations exist at the N position and two variations exist at the Y position)
This short primer mixture would have a degeneracy of eight degrees
It is recommended that primer degeneracy not exceed 100-200 fold degeneracy
If possible avoid placing degenerate bases near the 3' end of the primer since correct binding is crucial there
Inosine can be substituted at degenerate bases to reduce degeneacy. Inosine will usually bind to all possible bases, however it may result in loss in specificity and may form side-chain products with G bases. In general, it is best to limit the number of inosines in a degenerate primer to three or four.

Receiving Primers and Making Working Stocks

1. Before you perform a PCR reaction you must have the appropriate primers for the reaction. Often this means designing and ordering the primers (see above). When new primers arrive in the lab there are a number of steps to go through before they can be used in a PCR reaction.
2. The first thing to do is look at the synthesis report that came with the primer. Next to the primer write the lab primer number (PR1, etc). (If you are unsure check the Oligo Primers notebook (found near the catalogs) to see what the last assigned number is and then add 1 to this number)
3. Also give the primer a title revealing its use and write this on the synthesis information sheet. For example, "To make a mutant Pax BS2 in lin-48".
4. Now make a photocopy of the synthesis information. One copy will be stored in the notebook for the project for which the primer is being used. The other copy is stored in the lab notebook labeled Oligo Primers (found near the catalogs).
5. Enter the primer into the data base (more on this later).
6. The primers come as very small pellets in tubes. The pellet needs to be re-suspended in order to be used. To tell what the volume will be for suspension you will need to look at the synthesis information for the primer:
You want to resuspend as a 100uM (100pmol/ul) solution
Therefore, on the synthesis report find the amount of primer in nmol and multiple this by 100 to find the resuspension volume (in ul)
Example: 14.6 n mol of PR1 was shipped. It will be resuspended in 146 ul TE.
To resuspend:
7. Spin down the primer tubes (7 second short spin).
8. Add the amount of TE needed for a 100uM concentration (see directions in step 6 if you are unsure).
9. Label this tube with the PR# and the concentration.
10. Tap the tube to help dissolve the pellet.
11. Dilute immediately to working stock.
To dilute to make a working stock (25pmol/ul):
12. Spin down the primer tubes (7 second short spin).
13. Get a sterile 1.5 ml tube for each primer and label it with the PR#, concentration (25pmol/ul), date, and your initials.
14. To each sterile tube add 30ul of TE.
15. To the appropriately labeled tubes, add 10 ul of each (100uM) primer stock.
16. Place the 100uM stock in the Primers Box in the freezer.
17. Working stocks (25pmol/ul) are used in PCR reactions and stored at -200C when not being used.

Buffers

NEB

Fermentas

Making PCR reactions

Thaw Reagents

1. Get wet ice.
2. Thaw template DNA, 10X dNTPs stock solution, and reaction primers on ice (~ 30 minutes).
3. Obtain one thick-walled, sterile PCR tube for each reaction (2nd drawer below centrifuge).
4. Label the tube according to your system for PCR reactions.
5. You will probably run a 25ul or 50ul reaction. The standard ingredients in such a reaction are as follows(Primer may vary based on the number being used and DNA and enzyme may vary based on concentration and activity):

Fermentas Reagents

25ul 50ul
15.25 ul sterile ddH2O 31.75 ul sterile ddH2O
2.5 ul 10X Buffer (w/(NH4)2SO4) 5.0 ul 10X Buffer (w/(NH4)2SO4)
2.5 ul 10X dNTPs 5.0 ul 10X dNTPs
1.5 ul 2.5 mM MgCl2 3.0 ul 2.5 mM MgCl2
1.0 ul 25 pmol/ul primer 2.0 ul 25 pmol/ul primer
1.0 ul Second primer (25 pmol/ul) 2.0 ul Second primer (25 pmol/ul)
1.0 ul template DNA 1.0 ul template DNA
+ 0.25 ul Taq polymerase + 0.25 ul Taq polymerase

NEB Reagents

25ul 50ul
16.75 ul sterile ddH2O 36.75 ul sterile ddH2O
2.5 ul 10X Taq Buffer (w/MgCl2) 5.0 ul 10X Taq Buffer (w/MgCl2)
2.5 ul 10X dNTPs 5.0 ul 10X dNTPs
1.0 ul 25 pmol/ul primer 1.0 ul 25 pmol/ul primer
1.0 ul Second primer (25 pmol/ul) 1.0 ul Second primer (25 pmol/ul)
1.0 ul template DNA 1.0 ul template DNA
+ 0.25 ul Taq polymerase + 0.25 ul Taq polymerase

Make a Pre-Mix (for multiple reactions with identical reagents)

1. Label a sterile 1.5ml tube "Pre-Mix".
2. If all reactions include the same primer(s) and/or template DNA include these in the Pre-Mix.
3. To determine volumes added to Pre-Mix take the number of reactions for which the mix will be used, add1, then multiply by the volume of the ingredient you would use for one reaction:

Ingredient Volume in Pre-Mix = (n +1) (Ingredient volume for 1 reaction)

(Example: Here is a recent PCR Mix made for six (NEB) reactions where template was identical):
35 ul 10X Buffer (w/Mg)
35 ul 10X dNTPs
245 ul sterile ddH2O
7 ul template
4. Aliquot the appropriate volume of Pre-Mix to each PCR reaction tube.
5. Add ingredients - except for Polymerase - that were not included in the Pre-Mix (primer, template, etc. to bring to full volume).
6. Tap to mix your reactions (50 ul, no enzyme).
7. Balance and spin down the reactions (7 second quick spin).
8. ENZYME: Taq Polymerase is added in a Hot Start after the first step of PCR. This technique is covered under Using the PCR Machine (below).

Using the PCR Machine (Hybaid PCR Sprint Temperature Cycling System)

1. Additional related information can be found in the Hybaid PCR Sprint Temperature Cycling System User Instruction Manual (Pages 17-26 are especially helpful).
2. Write your name and a time range for your reaction on the PCR Sign-up calendar.
3. Turn on the PCR machine and wait for it to warm up.

TO RUN A PROGRAM AS IT IS
1. In the Main Menu, with the cursor on Run, press ENTER.
2. Use the Ø keys to select the source directory of the program you want to run. Press ENTER.
3. Enter the program number using the Ø keys. The program name will appear (if assigned). Press ENTER to select.
4. Select the following defaults by pressing ENTER for each:
Hot Lid: AUTO
Loading Alarm: OFF
End Run Alarm: OFF)
5. At the prompt "Lid on During Final Step?" answer NO and press ENTER.
6. Select default control mode SIM TUBE and press ENTER.
7. Enter the number of samples and the sample volume by using the number keys (you must enter preceding zeroes, i.e., 050 for "50"). Press ENTER to proceed.
(Cycling will commence heated lid is selected as OFF. If heated lid is selected as ON then the lid will preheat first. If MANUAL is selected ENTER needs to be pressed before cycling starts. If AUTO is selected cycling starts automatically)

PERFORMING A HOT START (ADDING ENZYME)
1. When the first round of amplification starts (usually two minutes at 940C) retrieve your polymerase in a freezer cooler.
2. Set a P2 for 0.25 ul and put a barrier tip on it.
3. When time left in Stage 1 is under 10 seconds, press PAUSE.
4. Open the lid and add 0.25 ul of polymerase to each reactions (change tips each time).
5. Close and secure the lid.
6. Press CONTINUE to restart the cycling. (To check conditions during the run, see below)
WHILE A PROGRAM IS RUNNING ­ RUN SCREENS
1. While a program is running three separate run screens can be displayed. These screens reveal current conditions (temp, stage/step), temperatures range achieved during cycling, and estimated time remaining.
2. The different screens can be accessed by pressing the and Ø keys to scroll through each screen.

CONSIDERATIONS IN CHANGING A PROGRAM

There are a few reasons you might want to change certain aspects of a program. The aspect that changes most often is the temperature of the Annealing step (Stage 2, Step 2) To learn how to change a program ask someone for help or consult the Hybaid booklet.

Purification of PCR products (Qiagen)
(adapted from Qiagen protocol book page 19)

1. This protocol is adapted from the QIAquick Spin Handbook, page 19.
2. Open the QIAquick PCR Purification Kit (Catalog No. 28104) and check to see that the Buffer PE contains ethanol (checkmark on lid). (If not add 24 ml of distilled ethanol to 6 ml of the wash buffer PE)
3. Add 5 volumes of Buffer PB per 1 volume of PCR reaction. (Ex. 105 ul PB to 21 ul PCR reaction) Vortex.
4. Label a spin column and 2 ml collection tube for each reaction.
5. Pipet each Buffer/PCR sample to the corresponding labeled spin column.
6. Balance and spin down for 1 minute at max speed.
7. Discard flow through from the collection tube. (Do not rinse tubes!)
8. Return each spin column to the same collection tube. To wash, add 0.75 ml Buffer PE to each spin column.
9. Balance and spin down for 1 minute at max speed.
10. Discard flow through from the collection tube. (Do not rinse tubes!)
11. Return each spin column to the same collection tube, balance and spin down for an additional 1 minute at max speed.
12. Discard the flow through and put the collection tubes into a sharps box.
13. Add the spin columns to sterile, labeled 1.5 ml centrifuge tubes.
14. To elute the purified DNA into the new tubes, add 50 ul Buffer EB to the center of each spin column.
15. Let the columns stands for 1 minute.
16. Balance and spin down for 1 minute at max speed.
17. The resulting liquid in the labeled 1.5 ml centrifuge tube is your purified DNA. Store properly or use immediately for further protocols.
18. If you feel it is necessary or you are unsure of the prep verify DNA recovery by spec or gel.

Making LB and/or LB Antibiotic plates

Estimated Time: 2 hours Yield: 45 3" diameter plates per1L of LB

1. Make LB w/ agar (usually 2 L)

Recipe for 2 L LB w/agar

Tryptone 20 g
Yeast Extract 10 g
NaCl 10 g
5N NaOH 400 ul
Agar, Granulated 30 g

2. Add about 1800 ml of dH2O to a sterile 2000 ml flask.
3. Add a magnetic stir bar to the flask and place it on a large stirrer.
4. Add 20.0 g of Tryptone to the flask and turn on the stirrer.
5. Add 10.0 g of Yeast Extract to the flask.
6. Add 10.0 g of NaCl to the flask.
7. Add 400 ul of 5N NaOH to the flask.
8. Remove the magnetic stir bar and pour the solution into a clean 2000 ml graduated cylinder. Use dH2O to bring the liquid level to 2000 ml.
9. Get 2 sterile, 2000 ml flasks. Pour about 1000 ml of solution into each flask.
10. Add the magnetic stir bar to one of these flasks and place it back on the stirrer. Add 15.0 g of Granulated Agar to the flask and stir the solution for a few seconds. Remove the stir bar.
11. Form an aluminum foil lid over the top of the flask. Tape the foil into place with one or two pieces of autoclave tape.
12. Repeat steps 9-11 for the other 2000 ml flask of solution.
13. Autoclave the two flasks of LB w/agar (Liquid setting, 30 min).
14. Retrieve LB w/agar about 50 minutes later and let it cool until it reaches a temperature tolerable for pouring (NOTE: Do not cool so much that is begins to solidify - About 30 minutes cooling time should be sufficient).
NOTE: If you are making LB w/ANTIBIOTIC plates thaw the antibiotic while the LB is cooling. Remove specific antibiotic(s) from freezer (Antibiotics Box) and thaw 1, 1.5 ml tube (containing 1.0 ml of 50mg/ml solution) for each 1 L of LB antibiotic. Unless you use a partial amount you can thaw the antibiotic tube at RT.

15. Wipe down pouring area of bench thoroughly with 95% EtOH solution to sterilize.
16. Obtain sterile plates of 3" diameter. You will need about 45 per liter of LB. Group the plates in stacks of 5 (lids up). Set the bags aside for later use in storage.
17. Label flask(s) that will receive antibiotic.
ADD ANTIBIOTIC WHEN LB IS COOLED: Add 1.0 ml of 50 mg/ml antibiotic to the appropriately labeled 1.0 L of LB. Re-cover flask with foil. Swirl the flask to mix the antibiotic into the LB.
18. Put on a lab coat to prevent LB from dripping on you as you pour. Have paper towels on hand since it is likely you will have drips or solution running down the outside of the flask.
19. Wipe off the bottom of the flask with a paper towel if it has been sitting in water.
20. Light flame. Remove and set aside the foil lid from the flask to be poured. Sterilize flask mouth in flame.
21. Pour LB into plates (start at the bottom of the stack and work up, holding the remaining empty plates and lids in your non-pouring hand as you pour). Pour into each plate until you just see the bottom of the plate covered with LB, and then stop pouring.
(NOTE: If you set down the flask to do something (for instance, to clean up some LB on the bench) cover the flask with the foil lid and flame the mouth before you start pouring again.)
(NOTE: If there are many bubbles on a plate you can use your burner flame to pop them CAREFULLY)
22. Pour all of the LB. If the last plate is insufficiently filled discard it into the sharps/burn bag.
23. Repeat steps 18-22 for the other flask of LB. If you pour different types of plates, keep the groupings of plates separate and label them.
24. Let plates stand at RT for 24 hours before bagging and storing (NOTE: They may be used for experiments later the same day if needed).
25. Label each plate with a vertical marking on the side of its base following our color coding system:
LB plate = single vertical black band
LB carb plate = black band + red band
LB kan plate = black band + green band
LB cam plate = black band + blue band
26. Label storage bag with at least the date. Tape storage bag shut and store at 4C.

Pouring NGM (Nematode Growth Medium) plates

1. NOTE: You will need sterile filling tube/sinker/tip in order to pour the plates. If these items are not sterile, begin the sterilization process in advance of the steps below.
2. Make 4 X 2.5 L NGM agar (following the recipe under Solutions) in 4, 5 L carboys.
3. Autoclave on liquid cycle for 45 minutes.
4. Retrieve NGM agar (done 67-73 minutes after starting the autoclave) and let the carboys sit in the large 500C bath (Hill Lab ­ Room 932) for at lesat 50 minutes. This brings them to approximately 500C ­ cool enough to work with, but not so cool that it will solidify.
5. Wipe down the pouring area with 95% EtOH solution to sterilize.
6. Obtain an appropriate number of small (60 X 15mm) petri dishes from the Hill storage area (Room 932). You will need about 1000 small petri dishes for 10.0 L of NGM agar.
7. Use a razor blade to make an opening in the top a bag of sterile plates. Flip the bag and pull off the sheath so that 20 plates are stacked on the bench.
8. Starting at the top, pull off 5 plates at a time and flip them over onto the bench (you will get stacks of 5 facing up). Repeat steps 7 and 8 until you have sufficient plates ready for pouring.
9. Once the carboys have cooled sufficiently, add the following to each using sterile glass pipets and flame sterile technique (WARNING: Do not flame the cholesterol since it contains ethanol!):

62.5 ml 1 M PPB (pH 6.0) (if previously aliquoted this can be poured)
2.5 ml 1 M MgSO4
2.5 ml 1 M CaCl2
2.5 ml Cholesterol (5 mg/ml EtOH)
10. Swirl the carboys to uniformly distribure these ingredients throughout.
Set up the Wheaton Omnispense by doing the following:
11. Unwrap the dispensing tubing except for the wrapped ends labeled 'sinker' and 'tip'.
12. Unscrew and remove the (Heidolph) screen on the side of the Omnispense.
13. Unscrew and remove the grey clamp on the side of the Omnispense.
14. Wedge the tubing at about its midpoint into the appropriate space at the side of the Omnispense. Wind the red turning device until the tube is inserted (NOTE: Make sure the sinker end of tube is coming out of the bottom, and the tip end of the tube is coming out the top).
15. Plug in the Wheation Omnispense and turn it on by flipping the white power switch in the back left corner.
16. The settings should include: Manual (for foot pedal control), and 10.0 ml for volume dispensed. To scroll through the settings press the yellow button labeled 'sel'.
17. Make sure the foot pedal is properly connected to the Omnispense.
18. Position one of the carboys on a cart as close as possible to the Omnispense.
19. Carefully insert the sink into the first carboy. Keep things sterile and grip the tube through the foil.
20. Uncover the tip and you are ready to pour. Pour by stepping on the pedal once to dispense 10 ml into one plate. Work from the bottom up in your stacks of 5, using one hand to hold the tip and the other to hold the plates.
21. Try to avoid spills or overflow; they may cause contaminant growth. Also try to avoid swirling the plates too much as you move them. (This will cause agar to solidify on the lid of the plate)
22. If you touch the tip to anything (like the bench) just flame it to sterilize.
23. When you reach the bottom of a carboy, prop it up with something so that you can fill as many plates as possible. If your last plate is not completely filled throw it in the burn box. Move the second carboy close to the Omnispense and carefully move the sink into it.
24. Fill the empty carboy with hot water and let is sit. Dispense the NGM from the second, third, and fourth carboy into petri dishes.
25. When this is finished, fill up a flask with hot tap water. Place the sink into the flask. Place the tip into one of the carboys. Use the foot pedal to flush the hot water through the tubing.
26. Free the tube from the Omnispense. Wash the tube out in the sink. Rewrap it in foil as follows: the tip and the sink get wrapped individually, and labeled with autoclave tape. The whole tube including tip and sink are then further wrapped in foil with autoclave tape that is dated. The package is ready to be autoclaved so it can be used again (Wrapped Solids cycle).
27. Remove agar that remains in carboys and put it directly in a burn box.
28. Rinse the carboys thoroughly and set them to dry.
29. The plates need to sit for a day or two. Once they have setup bacteria can be applied to them.

Spreading Bacteria on NGM (Nematode Growth Medium) Plates

1. Plates are typically spread with E. coli about 1 day after they are poured.
2. Check the fridge for a bottle of LB +Op that is fully grown.
3. If there is a bottle then proceed. If not see Inoculating LB +Op.
4. Light flame.
5. Stick green plunger or Pipetman on the end of a sterile, 10ml glass pipet (but do not remove it from the sterile metal sleeve yet).
6. Open the bottle of LB +Op and flame the mouth and cap. Remove the pipet from the sterile metal sleeve and flame it.
7. Take somewhere between 10 and 12 ml LB +Op. Remove the pipet and tilt it at an angle to prevent dripping.
8. Flame the LB +Op bottle and cap. Screw the cap back on.
9. Add bacteria to plates as follows:
Work on stacks of 5 plates from the bottom to top. Use one hand to hold the plates and lids and the other one to add the bacteria via the pipet
As you finish a stack of plates, push it aside
Add just a drop or two of bacteria to each plate
Try not to add too much bacteria and try to keep the pool of bacteria mostly in the middle of the plate
Keep the pipet at an angle of 30-45 degrees to prevent dripping on the bench
10. When the pipet is empty, add it to the bleach cleansing solution to be stored until it is washed.
11. Repeat steps 5-10 over with a new pipet each time until all plates are spread or you run out of LB +Op. Each round of 10 ml should allow you to spread approximately 70-100 medium plates or 6-8 large plates.
(If you run out of LB +Op, check the fridge for more. If there is no more, you will have to inoculate more (see below). After you spread the last plate you will probably have some LB +Op left in the pipet. Squirt this into the burn box or wash down the drain. Put the pipet in the bleach cleansing solution)
12. About a day later get an empty bin and wipe out the inside using a paper towel and 95% EtOH. Place the plates in stacks of 10 in the bin. You should be able to get 23 stacks (230 plates) in one bin. Label the bin with "Plates Spread (date)".
13. Repeat step 12 until all plates are in labeled bins. If you run out of bins label leave left over plates to the side on the bench. Put down a piece of tape to label them. Add these plates to bins as they become available.

Mini Prep (Alkaline Lysis)

Theory: You start with a liquid culture of cells containing vector and plasmid, cosmid, or fosmid insert. Cells are lysed (P2), and then proteins and cell structures other than genomic DNA are removed. The reaction is stopped (P3) and a relatively purified DNA product is resuspended.

1. A liquid culture for each prep should be grown the night before you plan on doing the prep. You can vary the number of preps from each plate, but it is customary to do multiple cultures (in most cases at least 3 or 4). (NOTE: In some cases you may need more cultures ­ when you have blunt end cuts or cross compatible enzyme sites that may reorient in multiple ways during ligation, or when the bacterial plate containing the colonies is likely to contain different species).
2. Check for the following solutions: P1 w/RNAse (4C), P2 (RT), and P3 (4C). If you need to make any of these follow the directions in the Solutions section of the manual.
3. Get wet ice.
4. Get 3 sterile 250 ml flasks to aliquot the P1, P2, and P3. Label them respectively with tape. Use a pipet and Pipetman to aliquot the appropriate amount of each stock to its individual 250 ml flask (You will need 200 ul of each stock for each prep. If you have 24 preps you will need 4.8 ml of each. Always round up to have extra to account for pipet errors).
5. Put the P1 and P3 stocks back in the refrigerator.
6. Label a sterile 1.5 ml tube for each liquid culture that you intend to mini prep. The label should include the name of the culture/prep.
7. Transfer 1.5 ml of each liquid culture to its corresponding tube. Set a P1000 to 0.75 ml and pipet twice. Use a flame to sterilize the mouth of the culture tube between transfers. Switch tips between cultures!
8. Balance the 1.5 ml tubes in the centrifuge and spin for 2 minutes at maximum speed.
9. While tubes spin, get a 250 or 500 ml flask to use for liquid waste during the mini prep.
10. After the spin you should observe a sizable pellet in each of the tubes. If any of the tubes do not have a pellet or have a much smaller pellet make note of this in your notebook.
11. Pour off the supernatant from each tube into the liquid waste flask (Keep your eye out for pellets that are floating! You do not want to lose a pellet in the liquid waste).
12. After pouring off there will be about 100 ul of supernatant left in each tube. Use a P200 to pipet this off (into liquid waste, tip in sharps box). Be careful not to suck up pellet. Change tips between tubes.
(NOTE: Always in the case of cosmids (or if there were small or non-existent plasmid pellets in step 10) repeat steps 7-12 to collect more DNA.)

(NOTE: In the following steps varying volumes of P1, P2, P3 can be used. The important point is that equal amounts of buffer are used throughout. 200 ul works fine for most mi