SECTION 1 Lab Safety

In an emergency

Call Campus Heath Service 548-4848
Room NN256A

go directly to the
University Hospital Emergency Room

call 9-911

http://depts.washington.edu/hhpccweb/CampusHealth.html

http://www.ehs.washington.edu/forms/IncidentFillin.pdf

Tissue Culture Safety

1. Every lab member should attend the yearly Blood Borne Pathogen review class. http://www.ehs.washington.edu/forms/classes/bpecrform.htm

2. Everyone who works in the tissue culture room must:

Have taken or be taking the Hepatitis B vaccination (685-1071)

Sign a form with the occupational safety nurse declining the offer (548-6117).

3. No food or drink is ever allowed in the tissue culture room.

4. Gloves and lab coats should be worn when working in the tissue culture room.

5. If you are working with human tissue, you should also wear goggles.

6. Mouth pipetting is never allowed.

7. The “Biohazard Safety Manual” is located on the shelf over the bench by Peter’s office and in the K-089 tissue culture room. Familiarize yourself with its contents.

8. Contaminated sharps go into the red sharps biohazard container. This waste must be autoclaved before disposal.

9. Waste -- All tissue culture flasks must be treated with bleach for at least 10 minutes before disposal.

All waste in biohazard bags must be autoclaved before disposal.

NEEDLE STICKS --
Wash immediately with soap and lots of water.

Before 4:00 pm go to room NN 256A Campus Health Service

(548-4848).

After 4:00 pm go directly to the University Hospital Emergency Room

Fill out an Incident report http://www.ehs.washington.edu/forms/IncidentFillin.pdf

Pay attention to what you are doing when you work with needles and you won’t be stuck!!!!!

1. ALWAYS know the potential hazards of a chemical BEFORE you work with it. You can get this information from the MSDS (Material Safety Data Sheets). These are filed in alphabetical order in the MSDS book located above the bench by Peter’s office. You can obtain on-line MSDS information through the LSS system (see next page) or by calling 3-0467.

2. Check the Yellow Pages of the University Chemical Safety Manual on the shelf by Peter’s office for procedures concerning commonly used chemicals

3. Use the fume hood when working with toxic or carcinogenic chemicals.

4. Wear gloves, goggles, and a lab coat when working with chemicals.

5. Do not wear open toed shoes in the lab.

6. Mouth pipetting is never allowed.

7. Emergency eyewashbottles at every sink or use big hose on the sink.

8. Emergency shower– entry way K-081.

9. Spills – We have solutions to neutralize chemical spill on the chemical bench. One is for acids, one is for bases and one is for organic substances and toxins.

10. The “Chemical Safety Manual” is located on the shelf located above the bench by Peter’s office. Familiarize yourself with it’s contents.

11. Chemical inventory. Room and location in the front of the MSDS book list all of our chemicals and in the red binder on freezer 4. They are also listed on-line in the LSS.

12. LOG IN ALL INCOMING CHEMICALS ON THE INVENTORY SHEET ON THE
REFRIGERATOR.

Chemical control

Orders-- order the smallest reasonable amount of carcinogens or toxins. When possible, order the hazardous substance premixed so you do not have to weigh the powder.

Waste -- Hazardous waste disposal forms are in the front of the MSDS book. USE THEM.

Sink logs -- Federal and Washington State laws require a record of chemicals poured down the drain. Use the sink logs by the sinks to record your waste.

In an emergency

Call Campus Heath Service 548-4848 Room NN256A

go directly to the University Hospital Emergency Room

call 9-911

Chemical Spills

Copied from: http://www.ehs.washington.edu/Services/Spill_Response.htm

Chemical Spill Basics

Hazardous material spills that do not endanger workers in the immediate area may be cleaned up by area personnel who have been trained and are properly equipped to clean up the spilled material safely. Spill kits are available from University Stores during business hours.

Our spill kit is under the sink by the fumehood HSB K-081

The neutralizing absorbents are on the chemical bench in K-081

Hazardous material spills that cannot be safely adsorbed, neutralized, or otherwise controlled at the time of release by employees in the immediate release area are considered to be emergencies requiring outside assistance by the Seattle Fire Department (SFD), Environmental Health & Safety (EH&S), and possibly a spill cleanup contractor.

When in doubt about whether you need help or not, it is best to call for help. EH&S staff cannot clean up spills but can offer advice on how to handle spills. Call 206.543.0467.

When you need emergency help, do the following:

Alert others and evacuate all affected areas.
Pull the fire alarm if needed or report the incident to UW police:

UW Campus or UWMC dial 9-911

The UW Police will notify the Seattle Fire Department (SFD) who will respond, stabilize, and contain the spill. Environmental Health & Safety (206.543.0467) will advise SFD as needed. The incident may require use of a spill clean up contractor at the department's expense. All waste must be contained and labeled as instructed by EH&S.

Major Chemical Spills

Pull the Fire Alarm or call 9-911

A major chemical spill is:

One that has caused injury to personnel or is likely to cause injury, or
Uncontained and spreading out of the immediate area endangering other labs, or
Has the potential to cause a fire.

Pull the fire alarm if someone has been injured. This is the fastest way to get help and alert others nearby of the emergency. The Seattle Fire Department is the primary responder for major chemical spills.

Minor Chemical Spills

Call 206.543-0467

Laboratory employees are responsible for minor spills of the chemicals they commonly use. Cleanup of minor spills is part of managing your laboratory chemicals properly. EH&S can provide training and consultation but does not maintain a Hazardous Materials Response team.

If you can answer YES to the following 4 questions, it is safe for you to clean up the spill:

1. Do you know what chemical was spilled?

2. Do you know the hazards of the spilled chemical?

3. Do you have a chemical spill kit?

Can you protect yourself from these hazards?

If you answered NO to any of the above questions or need assistance with the spill cleanup, evacuate the area and call EH&S at 206.543.0467 for assistance. EH&S will help you find the answers to these questions or bring in an outside Hazardous Materials Contractor to do the cleanup for you.

Chemical Waste

Copied from: http://www.ehs.washington.edu/waste/wastechemical.htm

Routine Chemical Collection

We have a routine collection set up for 70% ethanol-30% water: Routine 2000

We have a routine collection set up for Xylene 99% ethanol 1% Routine 2022

Wastes that are generated on a regular basis may be set up as a routine collection. To qualify for a routine request pickup, the composition of the waste generated must be identical each time. This is an extremely efficient process for collection. Your routine waste is assigned a number that is kept in our database; when you call in or e-mail the routine number to be picked up, we automatically know the waste's composition, regulatory codes, disposal options, and where you are located.

To set up a routine collection you will need to fill out a Setup Routine Chemical Request Form.
If you already have a routine set up and would like to request a pickup, please fill out a Routine Chemical Collection Request Form or call us at 685-2848 with the routine number and quantity to be picked up.

Do I have a Hazardous Waste?

In general, chemicals that are caustic, corrosive, flammable, toxic, or explosive are considered hazardous. Be aware that not everything that is non-hazardous is non-regulated. Refer to the Chemical Waste Management Guide or call 685-2848 if uncertain.

Certain acids and bases may be neutralized and disposed of to the sewer by the person who generated the material. Specific instructions and neutralizing agents are available through EH&S at 685-3759.

Preparing Chemical Waste for Pickup and Disposal

Chemicals must be in an appropriate container compatible with the waste, with a securely fastened screw top lid. As a rule, containers that were designed for solid reagents should not be used for liquids.
All containers must be labeled with a UW Hazardous Waste Label (Acrobat Reader)), unless the container has the original manufacturer's label on it already. The label must state all components of the waste and their percentages (percentages must add up to 100%).
Complete a Chemical Collection Request Form and mail to Box 354400 or FAX to 685-2915. When filling out the form, do not use chemical abbreviations or empirical formulas. Be sure to list all components in the mixture using percentage by volume (percentages must equal 100%). You may list up to four containers of the same mixture per line on the form. Standard turn around time is 2-4 weeks. Be aware that large requests, volume, or number of containers may take longer to be collected.
If you have any questions, please contact the technologist collecting in your area at one of the following numbers.

Zone 3	HSB	685-2849
Zone 4	South Campus, HMC	616-3200

What if it's an unknown?

Unknowns present a serious problem for the University. Without an accurate chemical name, chemicals can neither be handled nor disposed of in a safe manner.

Unknowns should be processed for collection and disposal as soon as possible following discovery. Do not store these wastes in satellite areas. Any information, such as history and physical properties that can be provided to the hazardous waste staff will aid in the investigation and identification of unknowns.
Disposal companies will not accept unknown chemical waste without an analysis. To have an unknown scheduled for testing first complete a Chemical Collection Request Form and mail it to Box 354400 or FAX it to 685-2915. Please include a budget name and number on the form. Currently, the cost of analysis is approximately $83 for unknowns of less than 4L (1 gallon).
The problems presented by unknowns can be reduced by periodic examination and inventory of stock chemicals, promptly labeling new containers, and disposing of all unused and waste chemicals from a satellite area prior to a faculty or staff member's departure.

Instructions for LSS

1. To get into the system click on "Uwick” in the program files, then “SSH secure”.

2. Type curie.u.washington.edu for host name OR click on curie.u.washington.edu if it is an option.

3. Username= gollahon

Password= rablab1

5. When the tutorial screen appears, bypass by pressing return

6. At the next menu, chose Chemical Inventory Menu by choosing or typing “INV”

7. At the next menu, chose view, update, or insert

V= view

U= update (for changing, deleting, and looking at files)

I= insert (for inserting new chemicals)

9. Next screen will be blank version of screen that you will always be working with. To get to the files you want to look at or update, must enter building and room codes.

Bldg ID: HSK -- press return

Room ID: K 081 – press Shift return

10. The screen should fill with product information at this point if you are in the update mode.

11. While in update screen:

(.)= delete

(O)= save

(1)= see options

(4)= add a comment onto a product line

(7)= see the online MSDS

F3= gets you out of any screen to the previous screen

F2= help

for other commands, see the official LSS manual

TO ADD A CHEMICAL TO AN EXISTING FILE:

1. Log on, go to the INV menu and chose “I” = insert

2. Enter Bldg ID: HSK-- press return

Room ID: K 081-- press enter

3. Tab to the product name field

4. Type name of chemical and press enter on numeric keypad

5. Find the chemical from the list the will appear by using the down arrow key

6. Press enter to select the product and have it placed in the inventory

7. At the point, return acts like TAB to move between columns so that you can add info like “amount”, “unit,” “surplus,” etc.

8. In “unit” column, press enter to see available codes, use down arrow to select correct unit (e.g. ML) and press enter

*If Chemical is not in LSS, but you want to add it: at step #4 above:

1. Move cursor to blank row using down arrow

2. Type name of product, press return

3. Type “y” to confirm that you want to add, and press return

Radiation Safety

All laboratory members who use radioactive materials must take the Radiation Safety Course offered by Environmental Health and Safety

http://www.ehs.washington.edu/training/radclass.htm

http://www.ehs.washington.edu/forms/RSClass_reg_form.htm

Our laboratory is authorized to use ³H and ³²P. ³H has low energy (0.006 MeV) beta emissions and requires no shielding. The waste is regulated and must be disposed of in LSA boxes. ³²P has beta emission (0.7 MeV) and should be shielded with plastic.

The waste for ³²P is in SEPARATE boxes from the ³H.

All ³²P waste is in K-081 (main lab).

All ³H waste is in K-089 (tissue culture room).

Make sure you know where to put the waste for your isotope.

All liquid ³²P sewer waste MUST be recorded on the clipboard that is on the shelf above the PCR machines. At the present time, there is ³H in K-089 and ³²P in K-081.

The Radiation Safety Manual is on the shelf above the bench by Peter’s office.

NO FOOD OR DRINK IS ALLOWED IN ROOM K-089 or K-081. BOTH ROOMS MUST BE LOCKED WHEN NO ONE IS PRESENT. We can lose our license over these two things!!!

Performing Calibrations
http://www.ehs.washington.edu/RadSaf/Rad_Calibrations.htm

The UW Radiation Safety Office (RSO) operates an instrument calibration facility (call (206)543-6328 for more information). Costs of meter calibration at the UW facility are comparable to other calibration facilities.

It is not required that the RSO calibrate your instruments. Calibrations may be performed by any qualified agency or by the instrument owner, provided that it can be demonstrated that the calibration is performed correctly.

To insure compliance with state regulations:

All survey instruments must be calibrated at least annually.
A record of the results and the method of calibration must be kept on file in the lab.
The records must be made available to the RSO or the Washington State Department of Health.
Even if the RSO does not calibrate your instruments, it is important that you provide them with copies of your calibration records to ensure compliance with the UW's Radioactive Materials License.

Fire Safety

Fire Safety. You must leave the area when there is a fire alarm. Make sure your lab area is as safe as possible before leaving. Do not use elevators.

The fire extinguishers are located outside the lab door (room K-081) and in K-079 (flow cytometry room). You should know how to use them. There is a class for this if you would like to take it. http://www.ehs.washington.edu/forms/classes/fetform.htm

In case of a big fire, activate the fire alarm in the hall. Then leave the building.

Earthquake Safety

Class offered through EEHS http://www.ehs.washington.edu/forms/classes/edform.htm

When the Earth Shakes

If you're at WORK OR SCHOOL, DROP under a desk or sturdy table. Stay away from windows, bookcases, file cabinets, heavy mirrors, hanging plants, and other objects that could fall. Stay under COVER until the shaking stops. HOLD onto the desk or table. If it moves, move with it. Here are some additional tips for specific locations.
If you are in a HIGH-RISE BUILDING and not near a desk or table, move against an interior wall, and protect your head with your arms. Face away from windows. Do not use elevators. Do not be surprised if alarm or sprinkler systems come on.

After the Earthquake
At the University you should have a predetermined place for you and your co-workers to meet -- kiosk outside K-wing.

Evacuate slowly and carefully. Look before exiting to make sure there is no overhead danger.
As you evacuate take note of utilities. Look for wires arcing, water running, and the smell of natural gas.

· Report to the kiosk outside K-wing, and take note of who is missing and any injuries that may exist.

Call or send a runner to the Police on Boat Street to inform them of damage, missing persons, injuries, and utility damage.
At your meeting spot, assist others, and check on loved ones. Let someone know if you leave.

Hazard Hunt
Conduct a hazard hunt at work.. Most injuries occur from interior flying or falling items. Check at least the following items:

Top heavy free standing furniture, Heavy or breakable objects
Electronic equipment and appliances. Unsecured cupboard doors
Hazardous chemicals, Utilities (gas, water, electrical)

Needle Sticks

The UW Campus Health Service (CHS) is the program for occupational health needs of all UW employees, students, faculty, volunteers and other designated UW affiliates.

The CHS clinic is located at the UW Medical Center on the 2^nd floor (NN256A) next door to the Emergency Medicine Service or at Hall Health Primary Care Center.

Call (206) 548-4848

http://depts.washington.edu/hhpccweb/CampusHealth.html

http://www.ehs.washington.edu/Services/accinc.htm

In case of a needle stick.

1. Report the incident to Katy, Martin, or Peter.

2. Seek medical care in the CHS clinic or Emergency Room without delay.

3. Laboratory tests and medication for HIV post-exposure prophylaxis should be started
within 1 to 2 hours after exposure.

4. Fill out an incident report form http://www.ehs.washington.edu/forms/IncidentFillin.pdf

Problems with Cooling, Heating, the Building or Scientic Instruments

Chilled Water Failure in Flow Cytometry Room

Call Robert Davis at 5-9438

General Heating, Cooling, Fumehood, or Building Problems:

M-F call 3-3010 or put your request on line at http://www.washington.edu/admin/facserv/workrequest.html. You will need a budget number for any repair that is not heating or cooling. The air-conditioner in the flow room requires a budget # because it is not a standard part of the building.

Evenings and weekends call the University Police at 3-3010 wait for the end of the message and then dial 0. They will contact the physical plant manager who is on-call.

Autoclave Failure

Call Steris at 1-800-333-8828. You will need to give them the institution, the room number and the serial number of the autoclave. Account # 46860

Gravity Sterilizer                             Room K059        serial # 011589306
Vacumatic Sterilizer                        Room K059        serial # 011589305
Gravity Sterilizer (dirty autoclave) Room K092        serial # 012788315

Scientific Equipment Repair

   Pipetmen go to G-156 HSB along with a budget #
   Small equipment can be taken to T-287 along with a budget #
   Large equipment (eg. incubators) call 543-5580 again you will need a budget #

Laminar Flow Hood in TC

Kurt Geissel Email: kutis@u.washington.edu
Phone: direct (206) 685-9343 (with Voice Mail) department (206) 543-9510
Fax: (206) 616-3360

Telephone Problems

Call 3-0133

Hospital Laundry

Call Michelle at 206-521-1740 (for pick up call 3-6729)
Pick ups are on Fridays only

SECTION 2 Record Keeping, Animal Use
AND Computer Use

Lab Notebook

All experiments must be recorded in a laboratory notebook. Katy can supply you with a notebook.

1. Use a numbered bound notebook for all entries. Number the pages and use the first few pages for an index.

2. Use pen for all entries

3. Date all entries. Write the day and full date at the top of the page. Try to keep different experiments on different pages.

4. Record ALL experiments – Successes, failures, and things you don’t understand.

5. Enter primary data immediately. Do not keep notes on scraps of paper or paper towels to enter later. Record details -- amount, concentration, how solutions were made, time and temperature of incubations, centrifuge speed, mistakes you made along the way, etc. Make sketches or diagrams if necessary.

6. Explicitly list, label, and identify your controls or standards.

7. If you run your experiment on the flow cytometer, make a note of the protocol used and the file names. It is a good idea to make a copy of the list you give to Thong with sample numbers for your notebook.

8. End each record with your conclusion and plans. Do this as soon as possible. You should have some hypothesis of what you expected from the experiment. Did this experiment confirm your hypothesis or not?

Frozen Cell Lines Log

N₂ TANK LOG INSTRUCTIONS:

The frozen cell line catalog is in the TC lab K-089 (brown binder). In an effort to minimize database disasters, you should use this catalog and not the computer to look up, record, edit or remove entries. Judy and only Judy will edit the computer logs and update this catalog as needed.

THE FIRST SECTION IS AN ALPHABETICAL LISTING BY CELL NAME.

THE SECOND SECTION IS A RACK AND BOX LISTING.

INSTRUCTIONS FOR REMOVING OR EDITING:

1. Look up your cell line in the ALPHABETICAL LISTING.

2. If you remove a line, highlight it with the attached yellow pen.

3. If you edit a line, just write the appropriate change in pen next to the entry.

INSTRUCTIONS FOR ADDING:

1. Look up the appropriate slot in the RACK AND BOX LISTING.

2. Record your new entry in pen on the appropriate line.

3. Highlight the new entry with the attached yellow pen.

ParafFin block Log

Paraffin BLOCK LOG INSTRUCTIONS:

The paraffin block catalog is in the K-081 (brown binder) under the bench in the middle bay. In an effort to minimize database disasters, you should use this catalog and not the computer to look up, record, edit or remove entries. Jeanne and only Jeanne will edit the computer logs and update this catalog as needed.

THE LISTINGS ARE BY DRAWER AND ROW

INSTRUCTIONS FOR REMOVING A BLOCK:

1. Look up your block.

2. Make a slip of paper and label it with the name of the block you will be taking.

3. Highlight the block name with the attached yellow pen. Write your initials and date by the block name

4. Remove the block and place the slip of paper in the slot so we will know where the block belongs when you return it.

INSTRUCTIONS FOR ADDING:

5. Identify the slot and drawer where the block lives. There should be a piece of paper in the slot.

6. Remove the paper and replace the block

7. In the log write returned and the date.

Computer Use

Rabinovitch Lab Computers

The computers in the Rabinovitch Lab are maintained by the Department of Pathology. For computer support or questions go to the following web site. http://www.pathology.washington.edu/tech/ or call (206) 221-5790

Access. In order to have access to a computer in the lab you must be assigned a password. Peter or Mike will send a request to info@pathology.washington.edu along with the individual’s name and email address.

Computer glitches. Send a Tech request to Computer Support http://www.pathology.washington.edu/tech/

Computer Use. The computers in the lab are research tools and as such they are to be used to write papers and reports, process data, access journal articles, correspond with colleagues concerning research information etc. They are NOT for playing games or excessive correspondence with friends.

For questions concerning UW policy see the following web site:

http://www.washington.edu/computing/rules/

Improper use of UW computers and networks can get you into trouble. It is your responsibility to know the rules. These UW guidelines and examples of the rules and laws of the state of Washington will help you to use computing and networking resources appropriately.

Revised Code of Washington (RCW) - Laws passed by the State Legislature

RCW 9a.52.110 Computer trespass in the first degree.

RCW 42.52.180 Use of public resources for political campaigns.

RCW 42.17.260 Documents and indexes to be made public.

RCW 42.52.160 Use of persons, money, or property for private gain.

Washington Administrative Code (WAC) - Rules and regulations for all state agencies

WAC 292-110-010 Appropriate and inappropriate use of state resources. (revised 04/98)
[See UWeek article State changes guidelines on email use]

Executive Ethics Board

Frequently asked questions about email and Internet use

Washington State Attorney General's Office

Junk Email - Information from the Consumer Protection Division

Email and Computer Usage by Faculty and Staff - Notice from the Provost and Executive Vice President

When you establish a UW NetID you open a gateway to a wealth of computing resources at the UW and beyond. Remember that inappropriate use of these computing resources can result in loss of access to them.

Staff use of Uniform Access computers is subject to the approval of their departments and supervisors.

Electronic Mail (EMAIL)

UW-GS 5

(Revised) January 1998 http://www.washington.edu/admin/recmgt/uw.gs5.html

Electronic mail is a technology that allows for the written exchange of information in machine readable format. Email represents not the system, but the information communicated through the system. Email messages are public records when they are created or received in the transaction of public business. They must be retained as evidence of official policies, actions, decisions or transactions. Email messages are considered public record material with legally mandated retention requirements, and are subject to the same rules and regulations as those which govern the management of paper records. Email is managed by its content, not its format.

Purpose. Email is meant for informal correspondence and scholarly communications. It should not be used for official record-keeping purposes. (For further guidelines on the uses of email, see Knowing the Rules on the Computing and Networking page)

Electronic Management. The University of Washington does not have central processes or resources to manage email in a way that meets specific Washington State Code regulating the management of public records. Backup of folders may not exist or folders may only be kept for a very short duration, so inadvertent deletion of messages can result in loss of information.

Privacy. Confidential and sensitive information should not be sent via email. The privacy and integrity of an email message cannot be guaranteed. Also, once created, there is no guarantee that attempts to erase or delete email will be effective.

Release. Under the Public Records Act (RCW 42.17.250 et seq.), if requested by a member of the public, email must be transmitted to the UW Public Records Office for review and possible release. Tape or disk copies of deleted documents are also subject to the Public Records Act.

Litigation. Unless protected by legal privilege, email can and will be discoverable in litigation. This applies to email on disk or on a backup medium.

Legal Proceedings. Like other forms of records, and regardless of retention requirements, email pertaining to pending audits, or judicial or public disclosure proceedings must not be destroyed until the issue is resolved.

Email messages are subject to the guidelines in RCW 40.14 regulating the preservation and destruction of public records and as such are managed through records retention schedules.

Email that is considered to have no administrative, legal, fiscal, or archival requirements for its retention may be deleted as soon as it has served its reference purpose. Refer to UW-GS4 .

The following categories of messages have specific retention periods. Refer to the University General Records Retention Schedule or your Departmental Records Retention Schedule for the retention period of individual items.

These records must be printed out and saved as a paper document as it is difficult for a department or unit to retain electronic records since individuals, electronic media, and desktop hardware and software change. Backup procedures for desktop equipment are often neglected and disaster recovery routines are not practiced. The only way to assure the retention of information is to print it and file it by subject or function in the appropriate paper filing system.

Policy and Procedure Directives

Correspondence or memoranda related to official public business

Agendas and minutes of meetings

Documents related to legal or audit issues

Messages which document departmental/office actions, decisions, operations and responsibilities

Documents that initiate, authorize or complete a business transaction

Drafts of documents that are circulated for comment or approval

Final reports or recommendations

Appointment Calendars

Email distribution lists

Other messages sent or received that relate to the transaction of University business.

Animal Protocols

1. Any one who works with animals is required to attend the animal training session given by the Department of Comparative Medicine every year. See policy page: http://cer.hs.washington.edu/iacuc/policies/index.html

2. The booklet written by the Department of Comparative Medicine in on the lab safety shelf above the bench by Peter's office (Laboratory Animal Regulations).

3. You must have an approved animal protocol for any procedure you perform on animals. It is a good idea to list the number of the protocol used in your laboratory notebook when you do the procedure. Our approved protocols are in the back of the Laboratory Animal Regulations book. Protocol forms are available on-line http://cer.hs.washington.edu/iacuc/iacucforms/index.html

4. If you need help with a procedure see Katy; she has worked with animals for many years and is familiar with most animal handling techniques.

5. If you have questions about a protocol or procedure and Katy does not know the answer, veterinarians are on call 24 hr. a day to help. Call 543-6257 for assistance.

SECTION 3 TISSUE CULTURE

Tissue Culture Media Preparation

MCDB 1531( liter)

Bottle sterile water for irrigation

Autoclaved flasks

Bell filter

MCDB 153 from Sigma (in refrigerator crisper)

Sodium Bicarbonate (7.5% solution) Invitrogen/Gibco

· Pour about half of the distilled H₂0 into large beaker.

· Add MCDB 153 powder to the beaker.

· Rinse the MCDB 153 packet with bottle sterile distilled H₂0 and add it to
the beaker.

· Fill the beaker up to about 900 ml mark

· Stir the solution

· Add 15.7 ml of NaHCO₃

· While stirring, bring the final pH to 7.2 by adding appropriate 4N HCl or
4N NaOH.

· Using 1 liter cylinder, add enough distilled H₂0 to bring it to 1 liter
solution

· Filter with bell filter into 200 ml plastic flasks.

L-15 (1 liter)

Bottle sterile water for irrigation

Two autoclaved 500 ml bottles

Bell filter

L-15 from Sigma (in refrigerator crisper)

· Pour 1 liter of the distilled H₂0 into graduated cylinder.

· Pour about half of distilled H₂0 the large beaker.

· Add L-15 powder to the beaker.

· Rinse the L-15 packet with some distilled H₂0 and add it to the beaker.

· Fill the beaker with the rest of the water from the cylinder.

· Stir the solution

· While stirring, bring the final pH to 7.2 by adding appropriate 4N HCl or
4N NaOH.

· Filter with bell filter into 500 ml bottles.

Barrett's Media Preparation

Reagent	Final Conc.	1 liter	200 ml	Stock
MCDB-153	1 X	1 package
Hydrocortisone	0.4 mg/ml	80 ml	16 ml	50 mg/ml
EGF	20 ng/ml	2 ml	400 ml	10 mg/ml
Cholera Toxin	10^-10M	84 ml	16.8 ml	100 mg/ml
Adenine	20 mg/L	200 ml	40 ml	100 mg/ml
Bovine Pituitary Extract	140 mg/ml	140 mg	25 mg	[see below]
Fetal Bovine Serum	5%	50 ml	10 ml
Penicillin-Streptomycin	100 unit/ml	10 ml	2 ml	10,000 units/ml
Amphotericin B	0.25 mg/ml	1 ml	200ml	250 mg/ml
Insulin-transferrin- selenium	5mg/ml	1ml	200ml	10 mg/ml
L-glutamine	4 mM	20 ml	4 ml

Media storage: Store in refrigerator

--------------------------------------------------------------------------------------------

Preparation of Growth Factors

1. Hydrocortisone (5 mg/ml stock) Sigma H-0396

Dissolve in water and aliquot one ml vials and freeze

2. Epidermal growth factor R&D Systems Cat.# 236-EG

3. Adenine (100 mg/ml) Sigma A-2786

1 gram of adenine + 10 ml 1N NaOH

Aliquot in 1 ml

Store in freezer

4. Cholera toxin (100 mg/ml) Sigma C-8052

0.5 mg Cholera toxin + 5 ml dH₂0

Aliquot into microcentrifuge tubes

Store in the refrigerator

5. Bovine Pituitary Extract
**Preparation depends on source of BPE**

**Test new BPE before old lot # runs out**

Invitrogen/Gibco BRL catalog # 13028-014 (25 mg in 5 mls--store frozen)

-Thaw and place into centrifuge tube. Spin down 5 minutes

Add supernat to media

- Remove 5 ml of media and resuspend BPE pellet. Spin down
another 5 minutes. Add supernat to media.

- Discard pellet

6. Fungizone (Invitrogen/Gibco)

Thaw and aliquot into 1.7ml. microfuge tubes. Refreeze at -20.

7. L-glutamine

Aliquot in 8 ml volumes.

Store in freezer. Thaw until clear, then add to media.

8. Insulin-transferrin-selenium/ITS (10 mg/ml, Invitrogen/Gibco)

Biohazardous material.

50 mg ITS + 5 ml sterile dH₂0

Aliquot 200 mls into microcentrifuge tubes. Store in freezer.

Product Qty Price Company Product #

Adenine 1 7.80 Sigma A 2786

MCDB 153 1 69.00 Sigma M 7403

ITS 1 72.45 Sigma I 1884

EGF 1 120.00 R&D Systems 236-EG

Fungizone 1 10.80 Invitrogen/Gibco 15295-018

BPE(25 mg) 1 25.00 Invitrogen/Gibco 13028-014

Hydro-
cortisone 1 21.00 Sigma H-0396

Media is a modification of that described in Washington, K. et al. Gut 1994 35:879-883.

Counting cells

Ref. Selected Method in Cellular Immunology B. B. Mishell and S.M. Shiigi ed. W.H. Freeman and Co. pg15

1. Dilute cells in media (usually a total of 10 mls) and mix well.

2. Remove 10 ml from the tube sterilely.

3. Add the 10 ml cell suspension to one side of the hemacytometer.

4. Count the cells in area 1 through 4 (see figure below). Count only those cells touching tow of the outside borders.

5. Determine the cell number using the following calculation

Cell/ml = (average of number of cells per large square) x 10⁴ x 1/dilution
Ex. If you count 39 cells in four large squares.
Cells/ml = (39/4) x 10⁴ x 1/1 = 9.75 x 10⁴ cells/m
If you have 10 mls of cells than you have a total cell count of 9.75 x 10⁵

Calculation of Population doulbing

Population doublings or PDL are calculated in the following manner.

ln # of cells harvested

PDL= # of cells seeded______

ln 2

For example: You plated 1 x 10⁶ cells.

You harvested 4 x 10⁶ cells.

ln 4 x 10⁶

PDL= 1 x 10⁶ = 1.38 = 2 PDLs

ln 2 0.69

If your PDL at plating was 40.5, then your new PDL is 42.5.

Esophageal Squamous Cell Media

Reagent Final Conc. 500 mls Stock

KBM 1X

Bovine Pituitary 2 ml 13 mg/ml

Extract

Hydrocortisone 0.5 ml 0.5 mg/ml

Human Epidermal 0.5 ml 0.5 mg/ml

Growth Factor

Epinephrine 0.5 ml 0.5 mg/ml

Transferrin 0.5 ml 10 mg/ml

Insulin 0.5 ml 5 mg/ml

Triiodothyronine 0.5 ml 6.5 mg/ml

GA-1000 (Gentamicin, 0.5 ml 50 mg/ml,

Amphotericin B) 50 mg/ml

Calcium chloride .05 mM 83 ml 300 mM

L-Glutamine 20 mM 5 ml 200 mM

How to order ingredients:

Product Qty Price Company Product #

KBM w/o Calcium 1 47.00 Cambrex (Clonetics) CC-3104

BEGM SingleQuots 1 46.00 Cambrex (Clonetics) CC-4175

Calcium Chloride 1 15.00 Cambrex (Clonetics) CC-4202

How to use the BEGM SingleQuots kit:

1. Remove vial of retinoic acid and set aside in freezer. It will not be used.

2. Let remaining vials thaw overnight in refrigerator (preferable). Can also thaw at room temperature or in 37°C incubator, but must be added to KBM as soon as
they are thawed.

3. Wipe outside of vials with ethanol and let air-dry.

4. Carefully remove contents of each vial with either plugged Pasteur pipette or pipetman and add to KBM.

Warm media by leaving out at room temperature for about 30 mins. It is best to aliquot the amount you need before warming. Do not warm in water bath--this causes the media supplements to degrade faster. Media is good for approximately 2-3 weeks after addition of supplements. Media shelf-life can be extended by only warming the amount needed for each feeding. Repeated warming cause supplements to degrade.

Ref: Oda, D. et al. In Vitro Cell Dev. Biol. Anim. 1998 34:46-52

Feeding Cells

Ex. of Feeding Media: 500 mls DMEM

55 mls (10%) Fetal Bovine Serum or Fetalclone III
(Hyclone,SH30109.03)

5.5mls (1:100 dilution) of L-Glutamine

5.5mls (1:100 dilution) of Penicillin-Streptomycin

write on label “FBS” or”FC” to identify, and “modified” or
“complete”

There is a list of media for cell lines in the appendix and two notebooks with cell line guidelines in the lab.

To start a new flask of cells:

Non-Adherent lines (ex. Jurkats and HL-60) T-125 flask:

Take 5 mls of cells from old flask and place in new flask (label). Draw off most of remaining cells from old flask; leave 5 mls behind (3 mls if a Friday and cells won’t be fed until Monday). Dump leftover cells in bleach. To each flask add 15 mls prewarmed (37°C) RPMI + FBS. Screw caps all the way down, then turn back 1/2 turn. Put in CO₂ incubator.

To do a normal feeding:

Adherent lines (ex. HeLa, Barrett’s):

Take off all of media and place in bleach. Add prewarmed media (37 ⁰C), 15 mls for T75 flasks, 5 mls for T25 flasks. (See tissue culture book for volumes of other types of plates and flasks) If cells are confluent, they need to be split.

To split a flask: (T75, volumes change for different flasks)

1. Remove all old media and place in bleach.

2. Wash cells 1X with 3-10 mls of Versene buffer. Make sure entire flask gets rinsed and remove all versene.

3. Add 1X trypsin/EDTA (1ml/T25,2.5ml./T75 and 4-5/T150,T175) flask and place in incubator for 5-10 minutes. Do not let cells sit in trypsin longer than 10 minutes. Knock flask on bench to dislodge cells. Check under microscope to see that all cells have come off the flask.

4. To inactivate trypsin add media with FBS (at least 2x volume of trypsin), or straight serum. Add entire solution to 15 ml conical tube.

5. Count cells by pipetting up and down in tube and adding 10ml cell mixture to hemacytometer.

6. Spin cells in swing arm centrifuge at 1000 rpm for 8 min. ( If you are just using the cells to reseed flasks, you can skip this step if you are diluting the trypsin at least 1 to10 in media.)

7. Remove supernatant and resuspend pellet in media. At this point you can decide how many flasks to split the pellet into. Add equal volume of pellet plus media mixture to each flask and bring to final volume with more media. Write passage number, cell type and date on flask and place in incubator.

Freezing Cells

For adherent cells: 1) Trypsinize cells with 1X trypsin/EDTA

2) Inactivate trypsin w/ media + serum.
transfer to conical tube

3) Do cell count

4) Spin down for 8 min at 1000 rpms

For non-adherent cells: 1) Pull off media and transfer to conical tube

2) Be sure to leave 3-5 mls media if planning to
continue culture

3) Do cell count

4) Spin down for 6 min at 1000 rpms

Prepare freezing media:

40 mls supplement-free media

-use DMEM for cells cultured in DMEM, or
MEM

-use RPMI for cells cultured in RPMI,
Hanks, MCDB, or other non-DMEM media

5 mls FBS

5 mls DMSO

1. While cells spin collect cryovials; clearly label and date them

- optimum cell # per vial = 1 to 3 million/ml. for adherent cells,10 million/ml. for non adherent cells.

- e.g. 6 million cells harvested: use 3 vials with 2 million cells in each

2. After spin, pour off supernate, resuspend pellet in freezing media, and aliquot to cryovials;

-e.g. resuspend 6 million cells in 3ml. Freezing media and place 1 ml of cells in each cryovial = 2million cells/vial.

3. Load cryovials into Mr. Frostee and place in -80°C freezer ASAP; not good to keep cells sitting in DMSO at room temperature.

NOTE: when freezing large numbers of cells, Mr. Frostee can be temporarily placed in refrigerator freezer for up to 2 hours, but don’t forget to transfer them to the -80°C freezer!!

4 In at least 4 hours or as long as three days, Judy will move cells to N₂ tank, and record cryovial information in Blue Folder N₂ tank log.

5. NEVER TOUCH THE COMPUTER N₂TANK LOG!!

Thawing Frozen Cells

1. Remove from one to three vials from the liquid N2 freezer at a time, or place the vials on dry ice.

2. Warm the vials immediately by gently shaking in a 37 C water bath to thaw quickly.

3. When almost all of the ice has thawed, take the vial to the hood. Wipe the vials with 70% ethanol. Open the cryovial very slowly. Pipet the thawed cells very gently into the appropriate size flask.

4. Add media to the cells dropwise and gently tilt the flask to mix the cells and spread them over the growing surface. Be sure flask is labeled with cell name, passage or PDL, cell number, and date.

5. Incubate overnight to 24 hours or long enough for attached cells to lay down of suspension cells to settle. Change the media to fresh media.

Starting Primary Cultures from Barrett’s Esophagus Biopsies

**Remember to keep everything as sterile as possible. Biopsies should be considered as biohazardous material--be extremely careful when processing**

1. Collect biopsies in vial containing tissue culture medium without DMSO. Keep vial on ice until processing biopsies. Ideally, use around 6 to 12 biopsies from each patient.

2. Transfer biopsies to a sterile 100 mm tissue culture dish. Rinse at least twice with 10 mls of 2X antimycotic solution. Leave a little liquid in plate to prevent tissue from drying out while mincing.

3. Using sterile scalpel blades, mince biopsies until small enough to pass through the tip of a 10 ml pipette (make sure to pre-moisten the inside of the pipette with media to prevent tissue from sticking to pipette).

4. Add 5 mls of tissue culture medium to pre-aliquoted collagenase III (25 mg in 15 ml conical tube). Pipette up and down until powder is completely dissolved.

5. Add collagenase solution to plate containing minces. Using a pre-moistened 10 ml pipette, transfer collagenase solution containing minces to a sterile T25 flask. Place flask in incubator at 37°C and let tissue digest for at least 5 hours.

6. Transfer minces and collagenase solution to 15 ml conical tube. Pipette up and down vigorously to dissociate tissue. Some visible tissue pieces will remain.

7. Pellet cells and tissue in centrifuge. Remove supernate and add fresh medium (use 1 ml per dish or well). Transfer to sterile 35 mm or 6-well tissue culture dish and place in incubator.

8. Leave dish undisturbed for 48 hours—check to make sure there is sufficient culture medium to prevent tissue and cells from drying out but not so much that tissue pieces float. If needed, add a little more media during this time.

9. Change tissue culture media, being careful not to disturb any pieces of tissue that have adhered to the plate. Add 2 mls of fresh media. Change media twice weekly. Epithelial cells should have explanted out by 1 week.

Collagenase preparation: (Collagenase III stored at 4°C)

Make fresh before processing tissue. Weigh out appropriate amount of collagenase III. Transfer to tube. Add enough modified MCDB 153 media for a final concentration of 1 mg/ml. Cap tube and tilt tube up and down to dissolve.

Modified MCDB 153:

See Barrett’s media section

Product Qty Size Company Product Price

Collagenase type 3 1 1 g Worthington LS004182 $126.00

Ref: Palanca-Wessels, MCA. et al. Gastroentrerology 1998 114:295-304

Set up of Esophageal Squamous cultures

(original protocol from Dolphine Oda--slightly modified by CPW)

**Remember to keep everything as sterile as possible. Biopsies should be considered as biohazardous material--be extremely careful when processing**

1. Keep biopsies on ice until ready to process.

2. Transfer biopsies into 60 mm tissue culture dish using sterile forceps (dip in ethanol and flame).

3. Mince biopsies using sterile scalpel blades until the pieces can pass through the tip of a 10 ml pipette. [When testing this, make sure to wet the inside of the pipette with sterile liquid or else tissue will stick to the inside of the pipette.] Add about 5 mls of 2X antimycotic and transfer to 15 ml conical tube.

4. Let pieces settle to bottom of the tube then remove supernatant with pipette. Rinse biopsies at least 5 times with 2X antimycotic solution (use total of 50 mls) by adding approximately 10 mls of solution and letting the tissue pieces settle to the bottom of the tube. Remove supernatant after each rinse.

5. After last rinse, add 6 mls of supplemented KBM (CC-3203 from Cambrex) containing 24 mgs of dispase II. Transfer biopsies into T-25 flask. Let incubate at least 16-24 h at 37°C.

6. Pellet tissue and remove supernatant.

7. Add 1 ml supplemented KBM. Using sterile plugged Pasteur pipette, triturate tissue for about 2 minutes until the suspension contains mainly single cells. Add 1 ml of DMEM + 10 % FBS (or any other media containing FBS) to inactivate enzyme.

8. Pellet cells and remove supernatant.

9. Resuspend in supplemented KBM and plate, dividing between two T-75 flasks. Return to 37°C incubator.

10. Can spin cells down the next day and resuspend in new KBM and replate.

11. Feed three times weekly with supplemented KBM. Should see islands of “cobblestone-like” cells by 7-10 days post-plating.

Antimycotic solution: Thaw frozen Amphotericin B (250 mg/ml) and Pen-Strep (10,000 units Pen/10 mg Strep per ml) stocks. [For Amphotericin B preparation, see Barrett’s media section.] Prepare 100X stock by adding 2 mls Amphotericin B stock to 18 mls Penicillin-Streptomycin stock. Label and store at 4°C. Make fresh 2X antimycotic solution just before processing tissue. Thaw 100X stock. Add 1 ml stock to 49 mls sterile PBS.

Dispase II: Weigh out 24 mg aliquots of Dispase II into 15 ml conical tubes. Store at 4°C. Reconstitute by adding 6 mls of supplemented KBM just prior to use.

Supplemented KBM: See Esophageal Squamous media section.

Product Qty Size Company Product # Price

Dispase II 1 5g Roche (Boehringer-Mannheim) 165 859 27.00

Soft Agar Assay for Colony Formation

Note: All volumes are calculated to cater for 4 plates per point.

Base Agar

1. Melt 3 ml aliquots of 1% DNA grade agarose in heat block set to 100°C for about 1 hour until melted.

2. Reduce temperature of heat block to cool agarose to 40°C (about 1 hour). Meanwhile, also heat 2X MCDB 153 + 10% FBS to 40°C.

3. Mix equal volumes of the two solutions to give 0.5% agarose + 1X MCDB 153 + 5% FBS.

4. Add 1.5 mls of mix per 35 mm petri dish/well of 6-well plate. Allow to set. Plates can be stored at 4°C for up to 1 week.

Top Agar

1. Melt 3 ml aliquots of 0.7% DNA grade agarose in heat block set to 100°C for about 1 hour until melted.

2. Reduce temperature of heat block to cool agarose to 40°C (about 1 hour). Meanwhile, also heat 2X MCDB 153 + 10% FBS to 40°C.

3. Remove plates from 4°C about 30 minutes prior to plating to allow them to warm up to room temperature (or place in 37°C incubator). Label plates/wells with base agar appropriately.

4. Trypsinize and count cells. It is very important to have a positive control line (e.g. ras transformed).

5. Adjust cell count to 200,000 cells/ml. This will give a final concentration of 5,000 cells per plate/well.

6. Place 0.1 ml of cell solution in a tube. Add 3 mls of warm 2X media to tube.

7. Add 3 mls of the melted agarose to the 2X media and cells to give 0.35% agarose + 1X MCDB 153 + 5% FBS. Gently pipet up and down to suspend cells evenly. NOTE: only do one tube at a time so that agar does not set prematurely.

8. Add 1.5 mls of mix per 35 mm petri dish/well of 6-well plate (usually plate out in triplicate). Allow to set.

9. Add 2 mls of regular growth media on top of agarose layer. Incubate assay at 37°C in humidified incubator for 10-14 days.

10. Stain plates with 0.5 ml of 0.005% crystal violet for >1 hour. Count colonies using dissecting scope.

**For agarose, mix appropriate amount of powdered agarose in PBS. Autoclave. While warm, place 3 ml aliquots in sterile polypropylene tubes with caps.

Freezing Whole Tissue

1. Fill plastic cryomold with OCT media.

2. Carefully place tissue in OCT media and submerge tissue gently using a wooden stick. Try not to create bubbles in the media.

3. Fill plastic beaker halfway with isopentane. Carefully submerge bottom of beaker in liquid nitrogen for 2 minutes to cool the liquid.

4. Carefully slide cryomold containing tissue and OCT media into cold isopentane.

5. Submerge bottom of beaker in liquid nitrogen until OCT media hardens (turns white).

6. Store frozen section wrapped in foil and placed in plastic bag at 70ºC before sectioning with cryostat. (Sections done by Gown Lab)

Product Company Size Product # Cost

Tissue Tek II Baxter (ASF) 15 x 15 x 5 M7114-13 $10.90

Cryomold 10 x 10 x 5 M7144-12

25 x 10 x 5 M7144-11

OCT Media Baxter (ASP) M7148-4 $5.13

2-Methyl Butane UW Stores # 0013-546 $58.84

(Isopentane)

Protocol described by Marilyn Skeely (Allen Gown Lab, UW Dept. of Pathology)

Lymphocyte Preparation from
Whole Blood

1. Collect 20 ml blood (assume »1-1.5 X 10⁶ WBC/ml whole blood) in green top tube (sodium heparin). Keep at room temperature. Have donor sign consent form and log book.

2. Add 3 ml of Ficoll-Paque at room temperature to each of six 15-ml clear plastic centrifuge tubes.

3. Dilute the blood in each 50-ml tube with an equal volume of RPMI 1640 without FBS (or use Hank’s balanced salt solution) at room temperature. This will give you about 16 ml of diluted blood/50-ml tube.

4. Mix gently, and then carefully, down the side of the 15-ml tube, layer the diluted blood (5-7 ml) on top of the Ficoll-Paque.

5. Put tubes in centrifuge; speed should be set at zero. Start centrifuge and increase speed gradually to prevent disturbing the layer. Centrifuge for 40 min at 1200 rpm. Make sure the BRAKE IS OFF. You do not want an abrupt stop. You want a gradual start and a gradual stop.

6. After spin, you should see a clear separation of layers.

Plasma (yellowish)

Mononuclear cell layer

(buffy coat)

Ficoll-Paque

(clear)

RBC

7. Using a sterile, clear, plugged Pasteur pipette, go through the upper layer down to the buffy coat and collect as much of it as you can and as little of the other layers as possible. The buffy coat contains the mononuclear cells (lymphocytes). Put the cells into a new 15-ml tube.

8. Add RPMI to dilute each tube up to 13-ml line (sometimes this will mean adding only 3-4 ml).

9. Spin 10 min at 1200 rmp.

10. Carefully aspirate off supernatant. Gently loosen the pellet by tapping.

11. Resuspend in 10 ml RPMI for washing.

12. Spin 10 min at 800 rpm.

13. Aspirate carefully. Loosen the pellet.

14. Resuspend in 10 ml of RPMI

15. Determine cell count (use hemacytometer).

Preparation of Mouse Embryo Fibroblast Cell Strains

(adapted from Laura Hays/ Meuth Lab protocol) 4/14/2004

1. Euthanize mouse and cut into abdominal cavity.

2. Tear open uterus and yolk sac with pair of forceps, remove embryos and tear away amniotic sac/placenta. Cut off the head and place it in an eppendorf tube. Place the body in a sterile petri dish with PBS (from a bottle of sterile PBS) – the amount of PBS should be enough to cover the bottom of the dish. Repeat for all – labeling both the plate and tube for each pup.

3. Cut the body in half and remove the liver (this removes the majority of RBC which will be toxic to the cultures). Take the remaining pieces to new dish with 10 ml of 0.25% Trypsin/EDTA. Then, holding the remaining embryo with forceps, slice several times with razor blade (This is a sort of tenderizing step, and is better if the embryo is not cut into several pieces). Mince finely with a razor blade. Pipette up and down a few times with a wide bore pipette. Incubate 37° for 15 min. Do these steps in batches up to 6 embryos before the 1^st incubation. Then, start working on the next batch.

4. Add 5 ml. more Trypsin/EDTA; pipette up and down a few times and incubate 37° for 5 min. (This step can go a lot longer so don’t worry).

5. Filter through Falcon 2360 100 micron filter into a 50 ml conical tube containing 5ml medium. Spin 10 min. 1500 rpm. Remove s/n very carefully with pipette (i.e. don’t use an asprirator on a vacuum line). If the cells do not pellet well or the pellet dislodges, respin. Resuspend in media.

DMEM with 4.5g/L glucose BioWhittaker 12-614F with:

1X glutamine GIBCO 25030-081

                                2X NEAA         BioWhittaker 13-114E

1X Pen/Strep BioWhittaker 17-602E

10% Fetal Clone III Hyclone SH 30109.03

6. Pipette into T150 flask or other flask and put in incubator. Incubate 24 to 48 hours in a 37C incubator with 5% CO2 and 3% O2. Do not let cells grow to more than 70-80% confluency before splitting.

7. Trypsinize and count cells. Freeze 1-1.5 million cells in cryovials in freezing media with 10% DMSO. Label with cell name, number, date and PDL 0. Seed flasks at 0.5-1 million per T150 or 175 or 0.25 million cells per T75. Check daily so that the cells are not allowed to become confluent.

Genotyping:

Take a 1mm cube from the head. Freeze the rest of the head.
Add tail piece and 50 ul of 0.1M NaOH to PCR tube
Boil for 30 min (PCR machine).
Add 50 ul of 1 M Tris (ph7.6) to neutralize
Vortex; quick spin to settle debris.
Need 1ul template for PCR
Run standard hCAT PCR (or other PCR genotyping protocol) for genotyping

Fixation and embedding cultured cells in agarose for paraffin

1. For adherent cells trypsinize and suspend in media with serum. For non-adherent cells simply harvest.

2. Pellet cells by centrifugation and wash two times in PBS.

3.Gently resuspend pellet of washed cells in 0.5% paraformaldehyde in PBS. Incubate for 20 minutes at room temperature.

4.Wash cells two times in PBS.

5.Resuspend the pellet in 70% ethanol and incubate overnight at 4 °C.

6.NEXT DAY—wash cells two times in PBS and pellet by centrifugation in 1.6 ml microfuge tube.

7.Remove most of the PBS and mix the cells gently with a toothpick.

8.Warm the cells to about 70°C and add an equal volume of 3% agarose (melted and at about 80°C) (Sea Plaque GTG Agarose Cambrex (FMC BioProducts) Cat. No 50111 made in water). Gently mix the cells with the agarose using a toothpick to minimize bubbles.

9.Cool completely and remove agarose cell pellet

10. Wrap in tissue paper and place in a tissue mold. Put in 70% ethanol and store at 4°C until ready to embed.

11. Take the mold to Hisopathology lab (C-417) with a budget number and instructions. The turn around time for processing and cutting blocks is about 1 week.

Section 4 Maintenance and calibration of Incubators

Changing the CO₂Tanks

Before beginning make sure the manifold has changed to the spare tanks by turning the

switch with an arrow painted on it to the side of the spare tanks (The side with pressure in the tanks). Also inspect all three gauges for the correct CO₂ pressure (obviously one will be almost zero).

1. To change the CO₂ tanks first turn the valves on the used tanks to off. Loosen the pigtail at the tank. Some gas will be released however the whistle from the escaping gas should end in a matter of seconds. If it does not retighten the pigtail and close the valve more.

2. After disconnecting the manifold from the tank screw the safety cap on. Always place the safety cap on the tank before attempting to move it. If the tank is dropped the safety top will protect the valve from breaking and possibly causing an explosion.

3. Undo the chains confining the empty CO₂ tanks.

4. Carefully shuffle the empty tanks (watch your toes) out of the area under the manifold by rolling it back and forth across the floor.

5. Remove the tag that is in the pocket attached to the tank. Rip off the bottom portion that has “Full” printed on it. Reinsert the tag so that the word “Empty” faces outward.

6. Undo the chains on the replacement CO₂ tanks. Loosen the Safety Caps if very tight (Do not remove them).

7. Roll the new tanks under the manifold, where the used tanks were and remove the safety top.

8. Roll the old tanks to the wall where the new tanks were and chain them so the empty sign faces out.

9. The screw at the top of the tank should be covered with a piece of foil and marked “FULL”. Remove the foil.

10. Wrap the screw with plumbers tape (on the shelf in TC) in the same direction as the pigtail will tighten.

In order to ensure a tight joint between the pigtail and the CO₂ tank an O-ring must be placed at the seal. Look down the end of the pigtail. If no O-ring is present use one of the black ones provided with the tank (usually attached with a rubber band). Tighten the pigtail to the tank. Slowly open the valve if it sounds like gas is leaking tighten the valve until it is not. Repeat for second tank.
Inspect the gauges one more time to see they are now all reading a normal pressure.
Order new tanks (See stores catalogue for ordering instructions).

Testing CO₂ levels in the incubators

Testing the level of CO₂ in the incubator is accomplished using the Bacharach kit located to the right of the two incubators stacked upright. Before beginning calibrate the Fyrite cylinder to zero. Add Fyrite CO₂ indicator refill (#11-0057) if necessary.

Testing the CO₂ level of incubator 1 and 2 (Forma Scientific)

Place the translucent yellow tubing from the Bacharach kit over the black knob with a hole marked "sample". Place the other end with the black shaped head over the top of the Fyrite cylinder. Depress plunger with head and pump bulb 20 times. Do not depress plunger while Fyrite is inverted. Release plunger. Invert Fyrite until fluid fills the clear tube. Orient the Fyrite to it's original direction. Wait until bubbles have cleared to get an accurate CO₂ reading.

CO₂ levels of incubators 3 and 4 (Sheldon Man. Corp.)

The basic procedure remains the same for these incubators with one small variation. Push the translucent tube inside of the larger tube sticking out the top of incubators 3 and 4 to take the sample.

After finding the CO₂ level record it on the paper taped to the outside of the incubator. Also record the temperature, CO₂ setting, and water level, fill water with autoclaved deionized water if necessary. If there is any anomaly in the incubator's temperature or CO₂ seek assistance. A normal CO₂ level should be about 5%.

Section 5 Flow protocols For Apoptosis and Proliferative Survival

Hoechst 33342/SYTO 11 Protocol for Apoptosis

Ref. Syto – Poot, M. Gibson, LL, Singer, VL (1997) Detection of apoptosis in live cells by MitoTracker Red CMXRos and SYTO dye flow cytometry. Cytometry 27:358-364

PI – (really Ho/FDA/PI) Stohr M and Vogt-Schaden M. A new dual staining technique for simultaneous flow cytometric DNA analysis of living and dead cells IN Laerum, OD, Lindmo T. Thorud E. (eds) “ FlowCytometry “ Vol IV Bergen; Norrway Universitetsforlaget pp 96-99 (1979)

Hamori, E., Arndt-Jovin, D.J. Grimwade, B.G. and Jovin, T.M. Selection of viable cells with known DNA content. Cytometry 1:132-135. (1980).

This protocol determines the relative number of live (Hoechst 33342 positive/PI negative), early apoptotic (Hoechst 33342 positive/PI negative, SYTO 11 low), dead (PI positive) cells and debris signals.

1. Make the following stock solutions:

1 mM Hoechst 33342 (in distilled water (do NOT use PBS, since phosphates will precipitate the dye)

10 mM SYTO 11 dye (add 2 mL of the 5 mM stock solution from Molecular Probes, Inc. cat. # S-7573 to 1 mL distilled water)

1 mg/mL Propidium Iodide in distilled water.

The Hoechst 33342 and propidium iodide solutions keeps in the refrigerator in the dark for weeks; the SYTO 11 has to be diluted on the same day as being used.

2. Bring cells into suspension; preferably at a density of 0.2 to 0.5 million per mL.

3. Add sequentially per mL cell suspension:

10 mL Hoechst 33342

10 mL SYTO 11

5 mL Propidium iodide

4. Incubate at 37 °C for 30 minutes.

1. Analyze on the flow cytometer using the HO-PI-SYTO 11 protocol.

Hoechst 33342 from Molecular Probes cat # H-1399

SYTO 11 from Molecular Probes cat # S-7573

Propidium iodide from Molecular Probes cat # P-1304

Hoechst 33342/SYTO 11/CMXRosamine Protocol for Apoptosis

Ref. Syto – Poot, M. Gibson, LL, Singer, VL (1997) Detection of apoptosis in live cells by MitoTracker Red CMXRos and SYTO dye flow
cytometry. Cytometry 27:358-364

Hamori, E., Arndt-Jovin, D.J. Grimwade, B.G. and Jovin, T.M. Selection of viable cells with known DNA content. Cytometry 1:132-135. (1980).

This protocol determines the absolute number of live (Hoechst 33342 positive/SYTO 11 and CMXRos high) vs. early apoptotic (Hoechst 33342 positive/SYTO 11 and CMXRos low) cells and debris signals. In this protocol two dyes that are sensitive to early events during are used. CMXRos is sensitive to changes in mitochondrial membrane potential; SYTO 11 separates early apoptotic from “normal” cells by a not yet known mechanism. The reason for using both dyes at the time is that not all pathways to apoptosis involve changes in SYTO 11 fluorescence; lowered mitochondrial membrane potential appears to be a more general indicator of apoptosis.

1. Make the following solutions:

A. 1 mM Hoechst 33342 (see Solutions section)

-- Be sure to use distilled water (do NOT use PBS, since phosphates will precipitate the dye)

B. 10 mM SYTO 11 dye

-- Dilute stock solution by adding 2 mL of the 5 mM stock solution to 1 mL distilled water.

C. 20 mM MitoTracker Red CMXRos in DMSO

-- Dilute 1:10 of the 200mM stock solution (see Solutions section) in DMSO.

Solution Notes:

The Hoechst 33342 solution keeps in the refrigerator in the dark for weeks.

The MitoTracker Red CMXRos solution keeps in the freezer in the dark for weeks.

The SYTO 11 has to be diluted on the same day as being used.

2. Bring cells into suspension; preferably at a density of 0.2 to 0.5 million per mL.

3. Add sequentially per mL cell suspension:

10 mL Hoechst 33342

10 mL SYTO 11

1 mL CMXRosamine

4. Incubate at 37°C for 30 minutes.

5. Analyze on the flow cytometer using the HO_Syto11_CMXRos_Apoptosis protocol. Cells should be analyzed as soon after staining as possible.

Reagents used in this protocol and their sources:

Hoechst 33342 from CalBiochem cat # 382065

SYTO 11 from Molecular Probes cat # S-7573

MitoTracker® Red CMXRosamine from Molecular Probes cat # M-7512

Hoechst 33342/SYTO 11/PI Protocol for Apoptosis

Ref. Syto – Poot, M. Gibson, LL, Singer, VL (1997) Detection of apoptosis in live cells by MitoTracker Red CMXRos and SYTO dye flow cytometry. Cytometry 27:358-364

Hamori, E., Arndt-Jovin, D.J. Grimwade, B.G. and Jovin, T.M. Selection of viable cells with known DNA content. Cytometry 1:132-135. (1980).

1. Make the following solutions:

A. 1 mM Hoechst 33342 (see Solutions Section)

-- Be sure to use distilled water (do NOT use PBS, since phosphates will precipitate the dye)

B. 10 mM SYTO 11 dye

-- Dilute stock solution by adding 2 mL of the 5 mM stock solution to 1 mL distilled water.

C. 1 mg/mL Propidium iodide in distilled water.

Solution Notes:

The Hoechst 33342 and propidium iodide solutions keep in the refrigerator in the dark for weeks; the SYTO 11 has to be diluted on the same day as being used.

2. Bring cells into suspension; preferably at a density of 0.2 to 0.5 million per mL.

3. Add sequentially per mL cell suspension:

10 mL Hoechst 33342

10 mL SYTO 11

5 mL Propidium iodide

4. Incubate at 37 °C for 30 minutes.

5. Analyze on the flow cytometer using the HO_Syto11_PI_Apoptosis protocol.

Reagents used in this protocol and their sources:

Hoechst 33342 from CalBiochem cat # 382065

SYTO 11 from Molecular Probes cat # S-7573

Propidium iodide from Sigma cat # P 4170

Hoechst 33342/SYTO 11/PI/CEN Protocol
for Apoptosis

Ref. Syto – Poot, M. Gibson, LL, Singer, VL (1997) Detection of apoptosis in live cells by MitoTracker Red CMXRos and SYTO dye flow cytometry. Cytometry 27:358-364

Hamori, E., Arndt-Jovin, D.J. Grimwade, B.G. and Jovin, T.M. Selection of viable cells with known DNA content. Cytometry 1:132-135. (1980).

Poot M., Gollahon, K.A. Gollahon, P.S. Rabinovitch (1999) Werner syndrome lymphoblastoid cells are sensitive to camptothecin-induced apoptosis in S-phase, Human Genetics 104, 10-14.

This protocol determines the absolute number of live (Hoechst 33342 positive/PI negative, early apoptotic (Hoechst 33342 positive/PI negative, SYTO 11 low), dead (PI positive) cells and debris signals. Thus, one can follow cells through various stages of apoptosis and decay.

1. Make the following solutions:

A. 1 mM Hoechst 33342 (see Solutions Section) -- Be sure to use distilled water (do NOT use PBS, since phosphates will precipitate the dye)

B. 10 mM SYTO 11 dye -- Dilute stock solution by adding 2 mL of the 5 mM stock solution to 1 mL distilled water.

C. 1 mg/mL Propidium iodide in distilled water.

Solution Notes:

The Hoechst 33342 and propidium iodide solutions keep in the refrigerator in the dark for weeks; the SYTO 11 has to be diluted on the same day as being used.

2. Bring cells into suspension; preferably at a density of 0.2 to 0.5 million per mL.

3. Add sequentially per mL cell suspension:

10 mL Hoechst 33342 (Molecular Probes cat # H-1399)

10 mL SYTO 11 (Molecular Probes cat # S-7573)

5 mL propidium iodide (Molecular Probes cat # P-1304)

1 mL CEN (CEN Singlet Cytometry Control; Riese Enterprises, Inc. Grass Vly, CA)

4. Incubate at 37 °C for 30 minutes

5. Analyze on the flow cytometer using HO_Syto11_PI_Apoptosis protocol. Cells should be analyzed as soon after staining as possible.

Reagents used in this protocol and their sources:

Hoechst 33342 from CalBiochem cat # 382065

SYTO 11 from Molecular Probes cat # S-7573

Propidium iodide from Sigma cat # P 4170

CEN are Singlet chicken erythrocyte nuclei, Riese Enterprises, Inc. Grass Valley, CA

Processing data by the Hoechst 33342/SYTO 11/PI/CEN Protocol for Apoptosis

1. Open your data files with MPLUS (in case of difficulties with this step ask Mike Shen for assistance).

2. Click on GATE 2D (blue field on the bottom of the base page).

3. Select PMT2 and PMT PK (red options on the GATE 2D page). This will select the PMT2 data (in this assay intensities of Hoechst 33342 fluorescence signals) on the X-axis and the PMT2 PK (peak height of the Hoechst 33342 fluorescence signals) on the Y-axis. A picture similar to the one shown in the top panel of the figure should appear.

4. Draw a diagonal as shown in the figure. Single cells will appear above the diagonal, while clumps will appear below the diagonal. This is because clumps have a relatively lower peak height than do single particles of the same total fluorescence intensity.

5. Click “DONE”.

6. Select PMT 2 LOG and PMT 5 LOG. This selects the Log distribution of the Hoechst fluorescence signals of all cells on the X-axis and the Log distribution of the SYTO 11 and PI fluorescence signals of all cells on the Y-axis. The green fluorescence of SYTO 11 “bleeds” into the channel of the red PI fluorescence (PMT 5). In this way “normal” cells (high SYTO 11 and PI negative) appear in the middle of the screen, “apoptotic” cells (low SYTO 11 and PI negative) appear in the bottom half of the screen and “dead” cells (PI positive) appear on the top of the screen.

7. Draw the regions as indicated in the bottom panel of the figure and save the result by clicking on “WINPRINT”. This saves the image and the signal numbers within each frame.

8. After you are done with all your data files, you can exit MPLUS by clicking “exit” and use PRINT ALL to print all saved files.

Gating to exclude clumps

PMT2 (Hoechst or DAPI)

Gating of “Normal”, Apoptotic and Dead Cells

and Debris in the Hoechst/SYTO 11/PI/CEN Assay

PMT2 LOG (Hoechst 33342)

Hoechst 33342/MTG/CMXRosamine Protocol for Apoptosis

Ref. Poot, M. and Pierce, R.H. (1999) Detection of changes in mitochondrial function during apoptosis by simultaneous staining with multiple fluorescent dyes and correlated multiparameter flow cytometry. Cytometry 35; 311-317.

Poot M and Pierce RH. Analysis of mitochondria by flow cytometry. Methods Cell Biol 2001;64:117-28

Poot M Mulitparameter Analysis of Physiological Changes in Apoptosis Current Protocols in Cytometry (2000) 9.15.1-9-9.15.7.

This protocol determines the number of live (Hoechst 33342 positive and MTG/CMXRos high) vs. early apoptotic (Hoechst 33342 positive and MTG/CMXRos low) cells and debris signals. In this protocol, three dyes are used. Hoechst stains all cells that contain DNA and resolves cells according to cell cycle stage; CMXRos is sensitive to changes in mitochondrial membrane potential; MTG reports the level of mitochondrial protein in cells. The reason for using MTG and CMXRos simultaneously is that this combination gives a better resolution between apoptotic (compromised mitochondria) and normal cells.

1. Make the following solutions:

A. 1 mM Hoechst 33342 (see Solutions section) -- Be sure to use distilled water (do NOT use PBS, since phosphates will precipitate the dye)

B. 20 mM MitoTracker Green (MTG) in DMSO. -- Dilute1:10 of the 200 mM stock solution (see Solutions section) in DMSO.

C. 20 mM MitoTracker Red CMXRos in DMSO -- Dilute 1:10 of the 200mM stock solution (see Solutions section) in DMSO.

Solution Notes:

The Hoechst 33342 solution keeps in the refrigerator in the dark for weeks.

The MitoTracker Red CMXRos solution keeps in the freezer in the dark for weeks.

The MitoTracker Green (MTG) solution keeps in the freezer in the dark for weeks.

2. Bring cells into suspension; preferably at a density of 0.2 to 0.5 million per mL.

3. Add sequentially per mL cell suspension:

10 mL Hoechst 33342

1 mL MTG

1 mL CMXRosamine

4. Incubate at 37˚C for 30 minutes

5. Analyze on the flow cytometer using the HO_MTG_CmxRos protocol. Cells should be analyzed as soon after staining as possible.

Reagents used in this protocol and their sources:

Hoechst 33342 from CalBiochem cat # 382065

MitoTracker® Green MTG from Molecular Probes cat # M-7514

MitoTracker® Red CMXRosamine from Molecular Probes cat # M-7512

Proliferative Survival Protocol

Poot M, Silber JR, Rabinovitch PS.A novel flow cytometric technique for drug cytotoxicity gives results comparable to colony-forming assays. Cytometry  2002 May 1;48(1):1-5

This procedure is based on the BrdU/Hoechst method according to:

Kubbies M and Rabinovitch PS (1983) Flow cytometric analysis of factors which influence the BrdUrd-Hoechst quenching effect in cultivated human fibroblasts and lymphocytes. Cytometry 3:276-281

Rabinovitch PS, Kubbies M, Chen YC, Schindler D, Hoehn H (1988) BrdU-Hoechst flow cytometry: a unique tool for quantitative cell cycle analysis. Exp Cell Res 174:309-318.

This protocol determines the number of proliferating and non-proliferating cells in cell cultures. The protocol involves exposure of cells to 5-bromodeoxyuridine, harvesting them, staining with a buffer that contains Hoechst 33258 dye and a known number of chicken erythrocyte nuclei (CEN). Incorporation of BrdU into the DNA leads to quenching of Hoechst fluorescence. In this way, cells that have undergone one or two rounds of DNA replication can be distinguished from resting cells. To resolve cells in the G1, S and G2 phase of the cell cycle samples are counterstained with ethidium bromide. By dividing the number of cells in the G1, the S and the G2 compartments of the first, second and third cell cycle by the number of CEN their numbers per unit of sample volume is determined.

1. Make the following stock solutions:

10 mM 5-bromodeoxyuridine (BrdU; Sigma B-5002) in phosphate buffered saline (PBS). You may need to warm the PBS to 37^oC, since 10 mM is close to the maximum solubility of BrdU. Filter sterilize and store at 4^oC in the dark. This solution keeps for at least one month.

Hoechst buffer (see next page)

200 mg/ml ethidium bromide. Dilute 200 mL of a 10 mg/ml stock (Sigma: E-1510) into 10 mL distilled water. Store at 4 ^oC in the dark. This solution keeps for at least one month.

2. Culture cells for at least one cell cycle duration in the presence of 100 mM BrdU. (See BrdU/Ho/EB protocol in this lab manual for BrdU treatment) Note: some cell types may be sensitive to BrdU, which leads to G2 phase arrest. In case an elevated level of G2 arrest is suspected, it is recommended to test a series of BrdU concentrations (see Rabinovitch et al., 1988). During BrdU labeling, all cell cultures should be protected from light by wrapping all plates or flasks in aluminum foil.

3. Harvest cells according to standard procedures (keep cells protected from light all stages of handling).

4. Strongly vortex chicken erythrocyte nuclei (named CEN Singlet Cytometry Control; Riese Enterprises, Inc. Grass Vly, CA) and add 1 mL of CEN per mL of Hoechst buffer.

5. Resuspend cells in Hoechst buffer; preferably at a density of 0.2 to 0.5 million per mL.

6. After at least 15 minutes of staining at room temperature in the dark (usually in a cabinet) add 20 mL of ethidium bromide per mL of cell suspension.

7. Stain for another 15 minutes in the dark.

8. Analyze samples on the flow cytometer by using the Z-Prol-Surv protocol.

Hoechst Buffer: 500 mls

Final concentration Amount to add
0.154 M NaCl 4.5 g NaCl

0.1 M Tris, pH 7.4 50 ml of 1 M Tris pH 7.4

0.1% NONIDENT P-40 500 ml of NONIDENT P-40

(IGEPAL CA-60)

1 mM CaCl₂ 1 ml of 500 mM CaCl₂

5 mM MgCl_{2 5
ml of 500 mM MgCl2}

0.2% BSA 1 g BSA

2.5 mg/ml Hoechst 33258 2 ml of 590 mg/ml Hoechst
33258

qs to 500 ml with dH₂0

Processing data for the Proliferative Survival Protocol

1. 1.Open your data files with MPLUS (in case of difficulties with this step ask Mike Shen for assistance).

2. 2.Click on GATE 2D (blue field on the bottom of the base page).

3. Select PMT2 and PMT PK (red options on the GATE 2D page). This will select the PMT2 data (in this assay intensities of Hoechst 33258 fluorescence signals) on the X-axis and the PMT2 PK (peak height of the Hoechst 33258 fluorescence signals) on the Y-axis. A picture similar to the one shown in the top panel of the figure should appear.

5. Click “DONE”.

6. Select PARAM A and PARAM B. This selects the distribution of the Hoechst fluorescence (quenched DNA) signals of gated cells on the X-axis and the distribution of the EB fluorescence (unquenched DNA) signals of gated cells on the Y-axis.

7. Click “Set Region” to draw gates of different cell cycle phases in various regions (see figure next page)

a. G0 cells in region #5

b. First cycle (G0+S+G2) cells in region #6

c. Second cycle (G1’ + S’ + G2’) cells in region #7

d. Third cycle (G1’’ + S’’ + G2’’) cells in region #8

e. CEN cells in region #9

Can include in gate #6

8. Regions from #5 to #9 are set in counter clockwise direction starting from lower right corner for region #5

9. Draw the regions as indicated in the bottom panel of the figure and save the result by clicking on “WINPRINT”. This saves the image and the signal numbers within each frame.

10. After you are done with all your data files, you can exit MPLUS by clicking “exit” and use PRINT ALL to print all saved files.

Calculations for Proliferative Survival

There is probably an easier way to describe this, but this is the way I understand.

A. List the gates in the following order then enter the number of events in each gate.

G0/G1 gate #5	1^st cycle Gate #6	2^nd cycle Gate #7	3^rd cycle Gate #8	CEN Gate #10
Enter # cells	Enter # cells	Enter # cells	Enter # cells	Enter # cells

B. Correct for the number of cells relative to CEN then multiply each corrected number by 10,000

Corrected # cells in G0/G1	Corrected # cells in 1^st cycle	Corrected # cells in 2^nd cycle	Corrected # cells in 3^rd cycle
(# cells gate #5/ # CEN) X 10,000	(# cells gate #6/ # CEN) X 10,000	(# cells gate #7/ # CEN) X 10,000	(# cells gate #8/ # CEN) X 10,000

C. Calculate the total number of proliferating cells

Add the Corrected # of cells in the 1^st cell cycle + (Corrected # of cells in the 2^nd cell cycle) divided by 2 + (Corrected # of cells in 3^rd cell cycle) divided by 4. This gives you the total # of proliferation cells (corrected for the number of divisions they have undergone). Then average the replicates.

D. Normalize the # of proliferating cells from each treatment group to the control. The control will always be 100%.

For example

Group	Total number of proliferating cells	Proliferative survival
Control	1276	(1276/1276) X 100 = 100%
2 nM CAM	1395	(1395/1276) X 100 = 109%
4 nM CAM	1024	(1024/1276) X 100 = 86%
8 nM CAM	259	(259/1276) X 100 = 19%

Section 6 Protocols for FISH and Antibody Stains

Slide preparation for sorts

1. To clean slides for sorts place pre-cleaned frosted slides into staining racks (holds about 20 slides).

2. Set up ethanol baths 30%, 70% and 100%.

3. Dip slides up and down three to five times in the 30% bath.

4. Dip slides up and down three to five times in the 70% bath.

5. Dip slides up and down three to five times in the 100% bath.

6. Allow the slides to air dry.

Deparaffinization of tissues on slides

Be sure to have tissue mounted onto CHARGED slides.

1. Bake slides in 90˚C heat block for 30 min.

2. Wash slides in Xylene 3x for 5 min each.

3. Dehydrate slides in 100%, 90%, 85%, 70% ethanol for 2 min each.

4. Dip slides in water and air dry.

5. Mark tissue area with diamond pen.

6. Store under nitrogen or at –20°C

Staining for Pericentrin

1. Wash slides with 1X Hanks buffered saline solution.

2. Cover with 80 mM PIPES for 1 minute. (PIPES was stored in refrigerator therefore 4°C).

3. Fix with methanol (-20C) for 5 minute. (Methanol stored in freezer therefore –20 C)

4. Hydrate with cold PBS until IF (stored in refrigerator if not stained immediately).

5. Incubate with PBS + 1% BSA + 0.1% Triton + 10% NGS (PBA buffer) for at least 15 min at RT. Do not let cells dry out at any point during the staining procedure.

6. Prepare affinity purified anti-pericentrin (M1-100) diluted 1:500 in PBA buffer. You will need 100ml per well (4 well slide).

7. Remove plastic chambers from slide. Cover each well with prepared antibody. Incubate in a humidified chamber for minimum of one hour at room temperature.

8. Prepare secondary GAR-FITC (Cappel) antibody as follows: Dilute antibody in PBA buffer 1:100 in an eppendorf tube. You will need 100ml per well plus 50 ml. Spin in microcentrifuge at highest speed for 10 minutes. Transfer to new tube leaving last 50 ml in bottom of tube (this removes any aggregates). Use only the antibody transferred to new tube—do not use remainder. Do not pipette up and down after spin. Leave on ice until needed.

9. After incubation with pericentrin antibody is done, wash slide with PBA buffer.

10. Cover each well with prepared secondary antibody. Incubate for at least 30 minutes light tight and in humidified chamber at room temperature.

11. Wash with PBA buffer

12. Counterstain with 1 part PBS to 3 parts PBS with DAPI (0.1mg/ml).

13. Wash with PBS.

14. Mount onto slide with DABCO. Some people edge the coverslip with fingernail polish for longer storage.

15. Store flat and light tight in refrigerator.

MPM-2/DAPI Protocol

Protocol from C. Sanchez in B. Reid Lab

1. Harvest both floating and adherent cells. Pellet cells and remove supernate.

2. Fix each sample in 500 ml of 1% paraformaldehyde in PBS for 10 minutes at room temperature.

3. Add 3 mls NST + 0.1% BSA. Count cells (optional). Centrifuge for 8 minutes @1500 rpm (463G). Remove supernate.

4. Resuspend in NST + 0.1% BSA at desired concentration. [Cells can be stored in refrigerator before continuing with the rest of the protocol.]

5. Transfer desired amount of cell suspension into eppendorf tubes. Add 1 ml of NST + 0.1% BSA + 10% NGS. Centrifuge 8 minutes same as above. Remove supernate.

6. Resuspend in 100 ml NST + 20% goat serum. Dilute MPM-2 antibody 1:20 in NST + 20% goat serum. Add 2.5 ml of diluted MPM-2 antibody to the tubes. Incubate on ice for 30 minutes.

7. Add 1 ml NST + 0.1% BSA. Centrifuge 8 minutes same as above. Remove supernate.

8. Resuspend in 100 ml NST. Add FITC-conjugated goat anti-mouse IgG (0.5 mg/ml). Incubate on ice for 30 minutes in dark.

9. Add 1 ml NST + 0.1% BSA. Centrifuge 8 minutes same as above. Remove supernate.

10. Resuspend in 200 ml DAPI. Run on cytometer.

NST buffer: solvent = dd H₂O, store at 4°

Compound	Concentration	Component for 500 ml
NaCl	146 mM	4.25 g
Tris base*	10 mM	0.6 g
CaCl₂	2 mM	2 ml
NONIDENT P-40	0.1 %	0.5 ml

*Adjust the pH to 7.4 with HCl after NaCl + Tris Base

Assay for Detection of Cyclin B by Flow Cytometry (small cell number)

1. Harvest and count cells. Centrifuge for 7 minutes at 1000 rpm (all subsequent centrifugation steps are similarly performed). Resuspend in PBS at 2 x 10⁶cells/ml and aliquot 250ml of each sample into a serum coated eppendorf tube (1.5 ml). Add 500 ml PBS and pellet cells.

2. Aspirate PBS. Resuspend in 250 ml of fresh PBS. Add 750 ml of paraformaldehyde and let cells fix for 10 minutes at room temperature.

3. Pellet cells. Aspirate supernate. Resuspend cells in 1 ml of 1% BSA, 0.25% triton X-100 in PBS and place on ice for 5 minutes.

4. Add 5 mls of 1% BSA in PBS. Pellet cells.

5. Aspirate PBS. Resuspend cells in 100 ml of PBS containing 1% BSA, 10% NGS. Add 10 ml anti-cyclin B antibody (BD (Pharmingen)) and incubate sample at 4°C overnight.

6. Add 500 ml 1% BSA in PBS. Pellet cells.

7. Aspirate supernate and resuspend in 250 ml DAPI. Run on flow cytometer.

Double Staining with Cyclin B and Pericentrin (sequential staining)

1. Incubate with PBS + 1% BSA + 0.1% Triton + 10% NGS (PBA buffer) for at least 15 min at RT. Do not let cells dry out.

2. Prepare anti-cyclin B (BD (Pharmingen)) as follows: Dilute antibody 1:1000 in PBA buffer in an eppendorf tube.

3. Incubate in anti-cyclin B (BD (Pharmingen)) in PBA buffer (at appropriate dilution) overnight at 4°C in a humidified chamber.

4. Wash with PBA buffer.

5. Incubate with affinity purified anti-pericentrin (M1-100) diluted 1:500 in PBA buffer at RT in a humidified chamber for minimum of one hour.

6. Wash with PBA buffer.

7. Prepare secondary GAR and GAM-antibodies as follows: Dilute antibodies in PBS + 10% NGS to appropriate concentration in an eppendorf tube. Spin in microcentrifuge tube at highest speed for 10 minutes. Transfer to new tube leaving last 50 ml in bottom of tube (this removes any aggregates). Use only the antibody transferred to new tube—do not use remainder. Do not pipette up and down after spin.

8. Incubate with secondary GAR-FITC (Cappel) 1:100 and GAM-Cy3 (Caltag) 1:80 in PBS + 10% NGS for at least 30 minutes light tight and in humidified chamber.

9. Wash with PBA buffer

10. Counterstain with 1 part PBS to 3 parts PBS with DAPI (0.1mg/ml).

11. Wash with PBS.

12. Mount onto slide with DABCO. Some people edge the coverslip with fingernail polish for longer storage.

13. Store flat and light tight in refrigerator.

Epi shake off procedure to isolate G1 cells for FISH and G2 cells for anaphase bridge scoring

Experiment: Isolation of epithelial cells using epi-shake off methodology

Date:

Solutions to make:

Buffer A: Hanks (without calcium or magnesium) + 5mM DTT.
Soaking solution: Buffer A + EDTA (To 4 mL of buffer A, add 40 mL 500mM EDTA). This solution should be heated to 37^oC.
Shaking solution: 4 mL buffer A + 40 mL of 500mM Ca²⁺ + 40 mL of 500 mM Mg²⁺ + 400 mL DMSO.

Method:

Remove colon sample from –80C freezer, pour contents of the sample tube into a petri dish, wash piece of tissue with PBS and pour out excess liquid.

Manipulate colon sample with the use of needles to bring the stroma side up (the sample should be concave). Put some superglue on a wooden stick, just enough to coat the bottom of the stick. With the stick, touch the stroma side of the colon sample and stick the rest of the sample upon the edge of the stick, manipulating it with an needle. Note: if the sample is less than 1mm x 3mm, DO NOT GLUE, put it directly into the soaking solution.

Stick a needle into the end of the stick, just as far enough so that the sample will be suspended in the soaking solution in the centrifuge tube.

Put the stick into the 15 mL centrifuge tube containing 2 mL SOAKING SOLUTION at 37^oC in a heat block. After 5 minutes, remove the stick from the soaking solution and remove the needle.

Place the stick into 2 mL SHAKING SOLUTION on ice and shake for approx 30 secs (this time will vary depending on exp been performed). You can embed the remaining tissue to check if epithelial cells remain after this procedure (this is optional).

The cells are now in Shaking solution. The cells can remain on ice in this solution or can be frozen at –80°C until ready to use. Take a cell count using 10 mL of the total cell suspension.

To proceed with the protocol, cells are centrifuged at 1,000 rpm for 8 min, cell pellet is washed with PBS, and centrifuged at 1,000 rpm for 8 min.

Resuspend pellet in 1% paraformaldehyde for 15 min at room temp. Centrifuge cells at 1,000 rpm for 8 min, remove supernatant and wash pellet in 1X PBS.

Centrifuge cells at 1,000 rpm for 8 min, remove supernatant and resuspend cells in PBS+1%BSA + 0.25% Triton-X-100, incubate for 5 min on ice.

Centrifuge cells at 1,000 rpm for 8 mins, remove supernatant and wash pellet in PBS, centrifuge cells at 1,000 rpm for 8 min, remove supernatant and resuspend pellet in DAPI (without NP40, but containing DMSO). At this stage the sample can be frozen -20C until ready to be flow sorted. The flow protocol to use to sort G1 and G2 cells simultaneously is named:

“JOS DAPICOMPENSATED 3-12-02”

Epi shake off: isolation of epithelial cells

updated by Rosana Risques, 10/16/03

(material needed is in a box labeled ‘epi’ over Judy’s bench)

Solutions to make:

Buffer A: Hanks (without calcium or magnesium) + 5mM DTT.
Soaking solution: Buffer A + EDTA (To 4 mL of buffer A, add 40 mL 500mM EDTA). This solution should be heated to 37^oC. Prepare 2ml per sample.
Shaking solution: 4 mL buffer A + 40 mL of 500mM Ca²⁺ + 40 mL of 500 mM Mg²⁺ + 400 mL DMSO. Prepare 2ml per each batch of epithelial cells collected (normally two).

Method:

Remove colon sample from –80C freezer, pour contents of the sample tube into a petri dish (wash with PBS is not necessary).

Manipulate colon sample with the use of needles to bring the stroma side up (the stroma is in the concave side and is shinny and brighter than the epithelial side, fig1). Put some superglue on a wooden stick, just enough to coat the bottom of the stick. With the stick, touch the stroma side of the colon sample and glue it. Try to fold up all the stroma surface towards the stick, manipulating it with an needle (fig.2). Note: if the sample is less than 1mm x 3mm, DO NOT GLUE, put it directly into the soaking solution.

Nail a needle into the end of the stick, just as far enough so that the sample will be suspended in the soaking solution in the centrifuge tube (fig 3).

Put the stick into the 15 mL centrifuge tube containing 2 mL SOAKING SOLUTION at 37^oC in a heat block or water bath. After 5 minutes, remove the stick from the soaking solution and remove the needle.

Place the stick into 2 mL SHAKING SOLUTION on ice and vortex high speed for approx 5 secs (first batch: superficial epithelial cells). If desired, take 10ul of cell suspension and mix with 200ul of DAPI (dil 1/20) to check in the microscope and estimate the amount of cells and the need to shake longer.

Place the stick into 2 mL SHAKING SOLUTION on ice and vortex high speed for approx 20 secs (second batch: deep crypt epithelial cells). (The cells can remain on ice in this solution or can be frozen at –80 until ready to use).

Centrifuge at 1,000 rpm for 8 min, remove supernatant and resuspend in 500-600 ul of DAPI with NP40 and DMSO (10 mg/ml) (in refrigerator 1). No extra washes are needed. Samples can be frozen at –20C until ready to be used.

Put the stick with the tissue in a new tube containing aprox. 2ml of 1XPBS. Vortex 2 min (third batch: mixture of epithelial and stroma cells). If desired these cells can also be centrifuged and resuspended in DAPI as the other two batches.

Put the stick with the remaining tissue (stroma) in a Petri dish. Scrape the tissue from the stick and add 1ml of DAPI. Mince the tissue using two scalpels. Then collect it in an eppendorf and filter it to a glass tube using a Millipore filter (SX0001300). Clean the filter after used with a toothbrush to recycle it.

Take the sample to be sort for G1 in the flow cytometer. The epithelial cells don’t need to be filtered, just pass through a syringe.

Fig 1. Fig2. Fig 3.

Tissue Digestion and CK Staining

(dissociating nuclei from formalin-fixed tissue) Updated by Rosana, 10/9/03

DAY 1

Cut 60uM sections, place them in a mesh bag and insert into green cassette with large holes. If tissue is large, put only one section per cassette. If small you can put up to 8 sections, cutting away the extra paraffin. If very small, wrap the tissue in special tissue paper or use tape to keep them on place. Usually Priscilla cuts them the day before and keeps them overnight in xylene.

Run samples through the automatic processor in Priscilla’s lab: (takes 2.5 hours)

Bin#1 100% Xylene
Bin#2 100% Xylene
Bin#3 100% Xylene
Bin#4 100% Xylene
Bin#5 100% ETOH
Bin#6 100% ETOH
Bin#7 95% ETOH
Bin#8 70% ETOH
Bin#9 50% ETOH
Bin#10 30% ETOH
Bin#11 deionized water
Bin#12 deionized water

Fill Bins#11,12 with water before beginning

Make sure other bins are full as well

With green cassettes in and covered, press “go to 1”

Push “lower”, “reset”, “autostart”

When finished (black dots don’t blink), press “raise”

Empty Bins#11,12 and wipe the water

Xylene removes paraffin, the ethanol gradient helps to rehydrate the tissue. If we don’t start immediately we can keep the cassettes in a beaker in water and in the fridge.

PREPARE:

collagenase 1mL aliquots (two per sample): remove from freezer 30 min before samples come out of processor and thaw in 37°C water bath with agitation – in glass beaker filled with water.

15ml FCS-coated falcon tubes (see note at the end)

Pour 2mL collagenase (two aliquots) into each falcon tube,

Remove samples from cassettes with tweezers and place the mesh bag into the collagenase.

Incubate in 37°C water bath with agitation for 4 hours. (Samples can be stored at 4°C after removing collagenase and filling the tube with 1XPBS).

Collagenase breaks cells apart. FCS prevents the cells to stick in the plastic.

PREPARE:

pepsin (one aliquot per sample, two if the tissue is in tape): remove from freezer (-80C at BB246) at least 15 min before samples are done incubating and thaw in 37°C water bath with agitation (in glass jar). Shake and vortex to mix before using

Triton-PBA, PBA (PBA is 1 gram of BSA per liter of PBS) and 1X PBS: place then on ice. They are in the fridge.

15ml FCS-coated falcon tubes (two per sample) and 1.5ml FCS-coated eppendorf tubes (two per sample: neg and pos)

timer for 10.5 min.

Add pepsin (one aliquot) to a 15ml FCS falcon tube.

Remove samples from water bath (one at a time), take mesh bag out of the tube, scrape the tissue using a plastic pipet and put it in the pepsin. If the tissue is in tape, take out collagenase with a platic pipet and add pepsin in the same tube.

Incubate in 37°C water bath with agitation EXACTLY 10.5 min. (Leave 2 min between samples at least).

Vortex 15 sec high speed.

Fill tube to top with cold 1xPBS to stop pepsin action.

Filter using 74 micron mesh (in drawer with scalpels) in new Falcon tube opening like a funnel (fold the mesh in 4 equal parts, poke sealed corner into tube). Take the sample with a plastic pipet, be careful not to overflow, squeeze mesh at the end. Fill to top with PBS for balance. Wait in here to finish all the samples

Spin for 10 min @ 1700rpm @ 2°C .

Remove supernatant (pour down sink, using orange pad at end to not lose pellet).

Add 500ul PBS and mix with pipet.

Count with hemocytometer: put 10ul in each chamber. Microscope 40X. To count we need min 25 cells/square.

10000/count = number of cells in positive tube

put the remaining in negative tube (if we don’t count or there are few cells, put 100ul in neg and the rest in pos)

Pepsin reverses the fixative action. The filtering is important to avoid clogging in the flow cytometer. The amount of antibody has been calculated for a specific amount of cells. Do not exceed it.

Add same volume of 0.2% triton-PBA to the volume of cells in both neg and pos tubes. Incubate on ice 3 min.

Add cold PBA to stop triton action. Use dropper, fill to top.

Spin for 10 min @ 1700rpm @ 2°C.

Pour supernatant into sink (using orange pad at end)

Triton makes the membrane of cells permeable, without destroying it.

Add antibody (in bottom drawer of fridge – neg. and pos.) and mix.

8ul of AE1/AE3 to positive samples

3ul of neg. antibody to negative samples

Incubate in the dark in the fridge overnight.

Anti-Cytokeratin (CAM 5.2)-FITC labeled, Becton Dickinson Catalog #347653 (positive)
Mouse IgG2a-FITC labeled, Becton Dickinson Catalog #349051 (negative control)
Anti-Rabbit IgG-RPE labeled (Fab)2, Sigma Catalog #P-8172 (RPE = R-Phycoerythrin) (negative control)
Anti-Cytokeratin (AE1/AE3) unlabelled, Roche Catalog #1-124-161 (no)
AE1/AE3 antibody needs to be conjugated Custom PE conjugation by Intergen at 1-800-468-7436 (positive)

FITC and PE are different fluorocroms. Both work equally well. Usually we use the PE ones. Both CAM5.2 and AE1/AE3 are antibodies for a broad spectra of cytokeratines.

DAY 2

Add PBA to fill tubes. Invert the tubes few times to clean.

Spin for 10 min @ 1700rpm @ 2°C.

Decant and re-suspend in 500ul working DAPI/DMSO (1:4 dilution pH 7.8, in fridge). If the sample is going to be analyzed the same day keep it in the fridge. For the next day keep it in the freezer.

Syringe with a 25G needle and transfer sample to a 3ml glass tube.

Sort in the cytometer for CK pos and CK neg cells. The sorted cells are in 1XPBS.

DAPI stains the DNA. It works instantaneatly, but the results look better if we wait 30 min before analysis.

NOTE: To coat the 15 ml and the 1.5 ml tubes with FCS just take 1ml of Fetal Calf Serum (FCS) and shake it up and down to cover all the surface of the tube (or pipet up and down in the 1.5 ml tubes). Re-use the 1ml of FCS up to 10-15 tubes.

Using a Counting Chamber

For microbiology, cell culture, and many applications that require use of suspensions of cells it is necessary to determine cell concentration. One can often determine cell density of a suspension spectrophotometrically, however that form of determination does not allow an assessment of cell viability, nor can one distinguish cell types. A device used for cell counting is called a counting chamber. The most widely used type of chamber is called a hemocytometer, since it was originally designed for performing blood cell counts.

To prepare the counting chamber the mirror-like polished surface is carefully cleaned with lens paper. The coverslip is also cleaned. Coverslips for counting chambers are specially made and are thicker than those for conventional microscopy, since they must be heavy enough to overcome the surface tension of a drop of liquid. The coverslip is placed over the counting surface prior to putting on the cell suspension. The suspension is introduced into one of the V-shaped wells with a pasteur or other type of pipet. The area under the coverslip fills by capillary action. Enough liquid should be introduced so that the mirrored surface is just covered. The charged counting chamber is then placed on the microscope stage and the counting grid is brought into focus at low power.

It is essential to be extremely careful with higher power objectives, since the counting chamber is much thicker than a conventional slide. The chamber or an objective lens may be damaged if the user is not not careful. One entire grid on standard hemacytometers with Neubauer rulings can be seen at 40x (4x objective). The main divisions separate the grid into 9 large squares (like a tic-tac-toe grid). Each square has a surface area of one square mm, and the depth of the chamber is 0.1 mm. Thus the entire counting grid lies under a volume of 0.9 mm-cubed.

Cell suspensions should be dilute enough so that the cells do not overlap each other on the grid, and should be uniformly distributed. To perform the count, determine the magnification needed to recognize the desired cell type. Now systematically count the cells in selected squares so that the total count is 100 cells or so (number of cells needed for a statistically significant count). For large cells this may mean counting the four large corner squares and the middle one. For a dense suspension of small cells you may wish to count the cells in the four 1/25 sq. mm corners plus the middle square in the central square. Always decide on a specific counting patter to avoid bias. For cells that overlap a ruling, count a cell as "in" if it overlaps the top or right ruling, and "out" if it overlaps the bottom or left ruling.

Here is how to determine a cell count using a standard hemocytometer. To get the final count in cells/ml, first divide the total count by 0.1 (chamber depth) then divide the result by the total surface area counted. For example if you counted 125 cells in each of the four large corner squares plus the middle, divide 125 by 0.1, then divide the result by 5 mm-squared, which is the total area counted (each large square is 1 mm-squared). 125/ 0.1 = 1250. 1250/5 = 250 cells/mm-cubed. There are 1000 mm-cubed per ml, so you calculate 250,000 cells/ml. Sometimes you will need to dilute a cell suspension to get the cell density low enough for counting. In that case you will need to multiply your final count by the dilution factor. For example, suppose that for counting we had to dilute a suspension of Chlamydomonas 10 fold. Suppose we obtained a final count of 250,000 cells/ml as above. Then the count in the original (undiluted) suspension is 10 x 250,000 which is 2,500,000 cells/ml.

Slide Fixation for CK-sorted cells dropped from Flow Cytometer

After the cells are sorted on to slides, and slides marked, and air dried overnight.
Next day proceed as follows.

1. Fix each slide in fume hood by adding 1 ml cold methacarn (3:1 methanol:acetic acid).

2. Let slides sit in hood for 8 minutes.

3. Remove methacarn by tilting slides onto a paper to towel to remove the methacarn.

4. Rinse the slides once in 2X SSC for 2 minutes.

5. Dehydrate in 70%, 85%, 90%, 100% EtOH, 3 minutes each.

6. Store the slides above liquid nitrogen.

Fixation of Sorted Cells for FISH

1. Allow the cells that have been sorted onto slides to air-dry overnight.

2. Mark the drop of cells on the bottom of the slide with a diamond top pen.

3. In the fume hood fix each sample by adding 1.0 ml of cold methacarn (3:1, methanol : glacial acetic acid) to the top of the slide.

4. Leave the slides at room tempurature in the fume hood for 10 min.

5. Remove the methacarn by tilting the slide and letting the liquid empty into a methacarn waste container.

6. Wash slides in 2X SSC for 2 min.; use fresh 2X SSC for each group of slides being washed.

7. Soak slides in 1% paraformaldehyde in PBS for 8 min. Only do this step if you have NOT already fixed cells in paraformaldehyde while prepping them.

8. Wash slides in 2X SSC for 2 min.; use fresh 2X SSC for each group of slides being washed.

9. Dehydrate the slides by soaking in ethanol at room temp. 3 min in each of the following concentrations: 70%, 85%, 90%, and 100% EtOH.

10. Store the slides under nitrogen vapor at -70° C.

FISH PROTOCOL – BAC PROBES

DAY 1

1. Take 35ml tube of formamide out of -20 freezer and place in 73°C water bath until thawed. When ready, add 2X SSC up to 50ml and pour into glass Coplin jar. Incubate at 73°C until solution equilibrates to 73°C.

2. Wash slides in 2X SSC for 2 minutes. (If slides have been stored overnight from cell prep, do this step. Otherwise, proceed straight from the cell prep).

3. Place slides in 70% ethanol @ room temperature for 2 minutes, then in 85%, 90% and 100% ethanol respectively, all at room temperature for 3 minutes each.

4. Let slides air dry for 3 minutes.

5. Prepare FISH probes: (these amounts are per slide)

BAC Probe

To BAC/Cot-1 Probe (400ng BAC and 20mg Cot-1) ADD :

1.0ml CEP Probe (Vysis)

8.0ml Trask Hybridization buffer OR Vysis LSI buffer

6. Denature probes at 73°C for 10 minutes then incubate at 37°C for 30-60 minutes before applying it to slides. The incubation at 37°C is very important! This is a pre-hybridization step.

7. Place slides in the formamide/SSC solution at 73°C in the water bath for 3 minutes.

8. Place slides into 70% ethanol at –20°C for 2 minutes, then in 85%, 90% and 100% ethanol respectively, all at –20°C for 3 minutes each.

9. Place slides on 37°C heat block for 2 minutes to let ethanol evaporate. While waiting, place paraffin around all of the ethanol containers because they can be used 5-10 times.

10. While slides are sitting on the 37°C heat block, add 10ul of the appropriate probe.

11. Cover Slip being careful to avoid air bubbles, this also to be done on the heat block.

12. Add rubber cement to edges of cover slip (using a disposable 3cc syringe) to seal,

and heat at 37°C for about 10 minutes until dry.

12. Incubate at 37°C overnight in moist chambers.

Day 2

1. Pre-heat 2 boats of 50% Formamide and 2 boats of 2X SSC to 45°C.

2. Remove cover slips with razor blade and place slides into gray holder.

3. Place slides in 50% Formamide at 45°C for 5 minutes. Agitating occasionally.

4. Repeat.

5. Wash slides in 2X SSC at 45°C for 5 minutes. Agitating occasionally.

6. Repeat.

7. Wash slides in 2X SSC + 0.005% tween for 2 minutes.

8. Add 100ml of Blocking Reagent and let sit 5 minutes.

9. Rinse quickly in 2X SSC + 0.005% tween.

10. Add 50ml of 11.7ug/ml of Avidin-Rhodamine solution I, cover slip and incubate at 37°C for 40 minutes.

11. Wash slides 3 x 3 minutes in 2X SSC + 0.005% tween at room temperature.

12. Add 100ml blocking reagent and let sit 5 minutes.

13. Rinse quickly in 2X SSC + 0.005% tween.

14. Add 50ml of 5ug/ml of Anti-Avidin solution, cover slip and incubate at 37°C for 40 minutes.

15. Wash slides 3 x 3 minutes in 2X SSC + 0.005% tween at room temperature.

16. Add 100ml blocking reagent and let sit 5 minutes.

17. Add 50ml of 11.7mg/ml Avidin-Rhodamine solution II, cover slip and incubate at 37°C for 40 minutes.

18. Wash 3 x 3 times in same buffer as in #13.

19. Apply counterstain – 13ml

20. Check slides on scope.

21. Store at –20°C under nitrogen.

Reagents

Avidin-Rhodamine solution I Anti-Avidin solution

5ml 1:6 Avidin-Rhodamine 10ml Anti-Avidin

45ml of Blocking Reagent to 1 ml of Blocking Reagent

(per slide)

Avidin-Rhodamine solution II Counterstain

5ml 1:6 Avidin-Rhodamine 5ml DAPI

45ml of 1X PBS 200ml Antifade

(per slide)

Digoxigenin Labeling of BAC DNA for FISH

10μl BAC DNA (~1μg)

32μl dH₂O

8μl DIG mix (Invitrogen mix)

1. Mix together in PCR tube

2. Incubate at 15°C for 3 hours

3. Put on ice and check size on gel (load 5 μl) – sample will look like a smear and should be 200-800bp

4. Add 5μl 0.5M EDTA to stop reaction

Preparation, dropping, and fixing of lymphocyte metaphase slides

Solutions to be made:

Hypotonic solution: 1.2 g potassium chloride,

0.4 g sodium citrate dehydrate

400 mL water

Carnoy’s fixative (methacarn): 3 parts absolute methanol

1 part glacial acetic acid

Method:

1. Place hypotonic solution in 37°C water bath to warm before use.

2. Centrifuge cells in suspension at 1000 rpm for 8 min.

3. Add 8 mL hypotonic solution, re-suspend well with Pasteur pipette.

4. Incubate for 20 min at 37°C.

5. Add 10-15 Pasteur pipette drops (about 0.5 mL) of fixative. Mix well.

6. Centrifuge at 1000 rpm for 8 min.

7. Aspirate supernatant, leaving 1 mL of solution. Flick tube to re-suspend well.

8. Gently add 3 pipettes of fixative, letting the fixative run down the side of the tube.

9. Re-suspend well with pipette.

10. Let sit for 10 min. at room temp.

11. Centrifuge at 1000 rpm for 8 min. (okay to refrigerate until ready to make slides).

12. Clean slides by placing in coplin jar containing 95% EtOH.

13. Rinse in water until water runs off in sheets, then shake off slide until only a film of water remains.

14. Pipette off supernatant from pelleted cells as close as possible without disturbing the pellet.

15. Add enough fixative to form a somewhat cloudy suspension (amount of fixative is dependent on the size of the pellet).

16. Set up dropping station.

17. Drop 15-20 μl onto each slide.

18. Allow slides to air-dry overnight.

Fixation (next day):

1. Mark cells with diamond pen to help locate during later experiments. If unable to locate cells, breathe on slide to help visualize drop.

2. Fix slides in methacarn for 8 min.

3. Wash slides in 2xSSC for 2 min.

4. Dehydrate slides in 70%, 85%, 90% and 100% EtOH at room temp for 3 min.

5. Freeze at -20°C under nitrogen.

FISH work on sorted cells

FISH Protocol for Direct Labeled Alpha-satellite Probes

(adapted from protocol by Shawna R. of T. Brentnall lab. Revised 8/31/00; revised again 11-18-02 by J. O’Sullivan)

17p FISH PROTOCOL

Solutions to make and/or heat before beginning protocol:

Formamide: Take 35 mL tube of formamide out of –20°C freezer and place in 37°C water bath until thawed. When ready, add 2 x SSC up to 50 mL and pour into Coplin glass jar. Incubate at 73°C until solution equilibrates to 73°C.

2 x SSC: Heat 40 mL of SSC in a Coplin jar at 37°C for at least 1 hour.

0.01N HCl: To 100 mL purified water, add 100 ml 10N HCl. Mix well and store at 4^oC for up to 3 months. Have a concentrated 10N HCl stock in the lab.

Pepsin working solution: To 40 mL 0.01N HCl, add 200 mL 1% pepsin stock solution. Aliquots of the 1% pepsin stock (stored in the –20°C freezer).

Note: thaw until no longer looks cloudy.

Day 1

1. Make 70% formamide: add 15 mL 2X SSC to 35 ml formamide, put Coplin jar (4 slides per Coplin jar)

2. Warm formamide (35 ml) in water bath at 73°C for 30 mins

3. Prepare FISH probes, vortex, spin and keep on ice until denatured

Prepare sufficient probe mixture for each slide and one extra per probe mixture for metaphase control slide

LSI p53 and CEP 17 Mixture Actual for 5 slides

Rack 7

Box 3

0.1 ml CEP 17 0.5 ml

1.0 ml LSI p53 5.0 ml

7.0 ml LSI hybrid buffer 35.0 ml

1.9 ml dH₂O 9.5 ml

4. Wash slides in 2XSSC for 30 min at 37C.

5. Incubate slides in formamide at 73°C 3 minutes.

6. Wash in 70%, 85%, 90%, 100% ETOH @ RT, 1 minute each.

7. Denature probes 73°C, 8 minutes, and then place in 45°C heat block.

8. Dry slides on 45 degree heat block ~ 2 minutes.

9. Add 11 ml probe while slides on heat block

10. Add cover slip immediately following each probe, 24x30mm

11. Add rubber cement (use 3 cc syringe)

12. Incubate slides overnight in moist chamber at 37°C, use slide moat with water.

Day 2

1. Heat 0.4 X SSC/0 .3% NP-40 to 73°C 30 minutes prior to adding slides.

2. Remove cover slips and place in gray slide holder.

3. Agitate slides 1-3 seconds and incubate slides at 73°C for 2 minutes.

4. Agitate 1-3 seconds and wash slides in 2X SSC/0.1% NP-40 1 minute.

5. Make anti-fade

100 ml anti-fade

2.5 ml 10 ng/ml DAPI (stored at –20°C)

6. Add 13 ml of anti-fade to each slide

7. Add cover slips

8. Store slides at –20°C under nitrogen in ziploc bags

Notes:

1. Replace EtOH series (both) after 5 sets of 4 slides (20 slides total)

2. Use LSI buffers NOT CEP buffers

3. Wash solutions should be pH 7

FISH Slide preparation

(1) For adherent cultured cells

1. Grow cells on a chamber slide.

2. Remove media and rinse once with PBS. Cover cells with freshly made methacarn (methanol:acetic acid 3:1). Fix for 8 minutes at room temperature.

3. Remove methacarn. Rinse once with 2XSSC.

4. Cover cells with 1% paraformaldehyde. Fix for 8 minutes at room temperature.

5. Remove paraformaldehyde. Rinse once with PBS and let air-dry overnight.

6. Remove plastic chamber after cells have dried on the slide. Make sure to scrape as much of the silicon gasket off as possible so a cover slip will fit evenly on the slide during hybridization. A razor blade helps with this. Mark where the cell are on the slide with a diamond tip pen.

(2) For cells in suspension:

1. Add 5 mM calcium/5 mM magnesium buffer up to 500 ml.

2. Add 10% NONIDENT P-40 (IGEPAL CA-60) for a final concentration of 0.1% (5 ml to 500 ml).

3. Centrifuge cells at 1000 rpm for 15 minutes.

4. Remove supernatant. Resuspend cells in 10-50 ml of deionized water.

5. Drop 8-10 ml onto each slide.

6. Let slides dry for at least 30 minutes at room temperature, then overnight if possible.

· Just make a couple of slides first and check them under the phase contrast scope to see if they are good. If the cells look crystalline (very bright fluorescent green color), stop here and try the cell prep again.

(3) Re-probing: Carefully remove coverslip and proceed with FISH—Day 1.

Probe Preparation:

Let probes come to room temperature before mixing. Centrifuge briefly to bring contents down to bottom of tube.

Probe mixes: (Use 5 ml of each probe mix per sample)

Alpha-satellite Chromosome 8 (Oncor)—FITC labeled

0.6 ml probe

30 ml hybridization buffer (Hybrisol VI)

*enough for 6 wells (4 well slide)

Alpha-satellite Chromosome 3 (Oncor)—TR labeled

0.5 ml probe

30 ml hybridization buffer (Hybrisol VI)

*enough for 6 wells (4 well slide)

Probes:

Cyclin D1/CEP 11 {rack 4, box 2}

Actual amount for 4 experimental & 1 control slide

2.25 mL dH₂O 11.25 mL

0.75 mL CyclinD1/CEP11 3.75 mL

7 mL LSI hybrid buffer 35.00 mL

Add in this order.

10 mL mixed probe per slide.

LSI p53 and CEP 17 mixture {rack 4, box 2}

Actual amount for 4 experimental & 1 control slide

1.9 mL dH₂O 9.5 mL

0.1 mL CEP 17 0.5 mL

1.0 mL LSI p 53 5.0 mL

7mL LSI hybrid buffer 35.0 mL

Add in this order.

10 ml mixed probe per slide.

Anti-fade reagent:

To 100 mL antifade reagent, add 2.5 mL 10 ng/mL DAPI (stored at –20^oC)

For 2-color FISH:

Heat each probe individually at 72°C for 5 minutes, then soak at 4°C until ready to use. Centrifuge briefly to bring contents to bottom of tube. Mix probes together prior to application. Apply 10 ml of combined probe mix to each well.

Notes on reagents:

· Formamide (Ultrapure): cat #75828 United States Biochemical (1-800-323-9750); 500 ml $46. Thaw and divide into 35 ml aliquots in 50 ml tubes. Store at -20°C.

· 20X SSC (Ultrapure): cat #15557-036 Invitrogen/Gibco; 4 L $62.00. Store at room temperature. Make up new 2X SSC for each experiment. 2X SSC with or without Tween 20 can be made up ahead of time and stored at room temperature.

· Ethanol washes: Ethanol can be reused up to 6 times. Store in paraffin covered containers either at room temperature or at -20°C.

· FISH probes: cat. # Varies, suffix for # either TR or FITC depending on label, ONCOR; $158. Comes with antifade.

After our meeting today and continued discuss with Alex, I propose the following designation for 4 reds/2 greens as normals for FISH. When scoring cells, and 2 sets of 2 duplets are seen, the cell will be scored as 4dd2; and, will be tallied as a 2/2 (similar to a 4DD green score for dicentrics being tallied as 2 green). The lower case "d" will minimize the confusion with "D" for dicentrics. For the Pancreas PO1 project, we can begin this designation for all future slides except for the concluding slides for the Bell study. For this, we have to wait for Alex's re-analysis of the Bell study.

As to what is a duplet, I propose that the 2 signals must be next to each other "as a duplet, or side-by-side". The distance between the two signals for a single duplet should not exceed the width of one signal. To be a normal 2 red, both duplets should meet the above within a single cell. At this time, a 3d2 is too controversial (thank goodness, low in frequency) to be taken out of the abnormal category. Wed 1/31/2001 2:44 PM Al Farrand -- Peter R and Teri B. concurred.

Karyotyping Adherent Cell Cultures
/Colcemid Treatment

(Protocol from UW Hospital Pathology Cytogenetics lab)

Harvesting the cells

1. Watch culture for mitotic cells. When culture is growing well, add 0.1 ml of colcemid solution (10 mg/ml) per 5 mls media. Depending on the proliferation rate of the cells, let the cells incubate in the colcemid from 2 to 12 hours. [Ideally, you would like to have a percentage of mitotic cells such that there will be 50 mitotic cells per slide. If colcemid is left too long, chromosomes will become shortened and it will be difficult to distinguish bands.]

2. Place hypotonic solution in 37°C water bath to warm while trypsinizing the cells. Collect the medium in 15 ml conical tubes. Trypsinize culture: rinse flask gently with versene (3 ml/T25; 4 ml/T75) and add rinse to the medium in the tubes. Add trypsin-EDTA to flask (1 ml/T25; 2.5 ml/T75) and let incubate @ 37°C for 5 minutes. Shake flask to detach cells. Tap flask on the counter (cells side down) several times. Check in microscope. Cells should be in a single cell suspension. Add cell suspension in trypsin to medium and Versene in centrifuge tubes. Spin 1000 rpm for 8 minutes.

3. Aspirate or pour medium to leave 1-2 ml in tube. Flick or pipette tube to resuspend cells well. Cells should be single cell suspension. Add 8 mls of 37°C hypotonic solution (see reagents below). Resuspend well with Pasteur pipette. Incubate 20 minutes @ 37°C. Add 10-15 Pasteur pipette drops (about 0.5 mls) of fixative (see reagents below). Mix. Spin at 1000 rpm for 10 minutes.

4. Aspirate medium to 1 ml. Flick tube to resuspend well. Gently add 3 pipettes of fixative by letting fixative run down the side of the tube. Resuspend well with pipette. Let sit 10 minutes @ room temperature. Spin 1000 g for 8 minutes. Can refrigerate until ready to make slides.

Slide Preparation

1. Clean slides by placing in Coplin jar containing 95% ETOH. Rinse in water until water runs off in sheets. Shake off slide till only a film of water remains. Place slides on wet paper towel.

2. Pipette off supernatant from pelleted cells as close to pellet as possible without disturbing the pelleting. Then add enough fixative to form a somewhat cloudy suspension (amount of fixative is dependent on size of the pellet.

3. Drop 1 drop of cell suspension with Pasteur pipette (can add more drops if cell concentration is low). Wait till drop spreads then add one drop of fixative with Pasteur pipette. Can use a kimwipe to wipe outside edge of slide if there is excess liquid. Let air dry.

4. Examine slides on low power (10X) phase contrast to visualize chromosomes.

Staining slides: G banding (done in Cytogenetics lab)

1. Bake slides at 95°C for 1 hour (if using cytogenetics oven, time using button #3 marked slides on timer on bench opposite from oven). This “ages” slides and decreases the trypsinization time.

2. Trypsinize slides for 30-60 seconds (length of time depends on cell type); use high trypsin concentration. While waiting, measure 4 mls of pH buffer into small beaker. When trypsinization is done, briefly rinse 2X through rinse buffer.

3. Lay slide face up on staining rack over sink. Add 1 ml of Wright stain to buffer in beaker and swirl to mix. Pour over slide in rack and stain 60 seconds (time is dependent on cell type). Rinse out beaker with distilled water. Rinse slide after staining is finished. Air dry.

4. To visualize, focus using 10X magnification on light microscope (use green filter). Add 1 drop of oil and use 63X objective.

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Reagents and Solutions

Colcemid (Invitrogen/Gibco #15210-016)

Hypotonic solution:

1.2g potassium chloride (KCl)

0.4g sodium citrate dihydrate (Na₃C₆H₅O₇.2H₂O)

in 400 mls ddH₂O

Carnoy’s fixative: (Make fresh)

3 parts absolute methanol : 1 part glacial acetic acid

Staining of Tissue OR CELLS WITH CATALASE (can be used for most antibody stains)

Materials:

Rabbit Anti-Human Catalase = in box “unlabeled primary antibody” in frig #2, top drawer
BSA (Bovine Albumin)= frig #1 in the white box on top shelve
Alexa 647 chicken anti-rabbit = frig #2, top drawer, 2^nd Antibody box
V=vertical in a coplin jar or bath, h=horizontal – the reagent is made up in a small quantity and applied directly to the slide.

Day 1

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If the slides are formalin fixed and cut from paraffin blocks:

Deparaffinization :

Be sure that slides are labeled in pencil and that the tissue is circled with a diamond pen (mark on bottom of the slides)
Transfer slides to a slide rack
Immerse Slides in Xylene 3x 5minutes, waste goes to glass bottle in hood (v)
Immerse slides in 100% ethanol 2x5minutes (v)
Immerse slides in 90% ethanol 1x5minutes (v)
Immerse slides in 85% ethanol 1x5minutes (v)
Immerse slides in 70% ethanol 1x5minutes (v)
Immerse slides in 50% ethanol 1x5minutes (v)
Immerse slides in 30% ethanol 1x5minutes (v)
Wash slides with ddH20 2x5 minutes with rocking (v)

Antigen retrieval:

Heat 10mM Sodium Citrate (pH 6.5) to 88°C in a water bath
Immerse Slides in Sodium Citrate for 15 minutes (v)
Allow to cool in 1xPBS (v)
MOVE STRAIGHT TO STAINING

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If the slides are fresh tissue culture chambers with living cells:

Fixation & Permeabilization:

Wash cells with PBS 2x
Fix for 2 minutes on ice with ice cold 10:1 methanol:acetic acid
Wash with PBS 2x
30’ at RT in 5% NGS + 0.005% Triton X-100 (blocking and permeabilization)
Remove the chambers

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Staining

Block 30’ at room temperature with 10% NGS in PBS (h)
Remove blocking solution by wicking
Apply primary antibody (1:100) (h-with coverslip) in PBS +3% BSA (optimize for your primary antibody)
Incubate overnight at 4°C in humidified chamber

Day 2

Remove coverslip and wash slides 3x5’ with PBS + 0.05% tween with rocking (v)
Apply secondary antibody (1:100) in 1% NGS +PBS (h)
Incubate at room temperature for 1 hour (in the dark from this step forward)
Wash 2x5’ with PBS+0.05% tween and 1x with PBS with rocking (v)
Remove slides from PBS, blot off excess liquid, add a drop of vectashield (can add DAPI to vectashield if nuclear counterstain is desired) and coverslip. Be sure to remove bubbles from under coverslip. Wipe off excess vectashield and coat the edges with nail polish.
Store in dark at 4°C until ready to view at microscope

RC DC Assay Protein Quantification

The RC DC assay is a colorimetric assay for protein quantification.

1. Prepare 3-5 dilutions of a protein standard from 0 ug/ml to 2 mg/ml protein. (using lysis buffer as the dilution buffer)

2. Pipet 25 ul of standards and samples into microfuge tubes.

3. Add 125 ul RC reagent I into each tube. Vortex, and incubate for 1 min at RT

4. Add 125 ul RC reagent II into each tube. Vortex, and centrifuge the tubes at 15,000xg for 3-5 mins

5. Discard supernatant by inverting tubes. Allow liquid to completely dry out from tubes (may leave overnight)

6. Add 127 ul Reagent A’ to each tube. Vortex, and incubate for 5 mins at RT.

5ul of DC reagent S to each 250 ul of DC reagent A = 255 ul Reagent A’

7. Add 1 ml of DC reagent B to each tube and vortex immediately. Incubate for 15 mins at RT

8. Pipet 200 ul of each sample into a 96-well plate. Triplicate for each sample.

Absorbance can be read at 750nm.

*BioRad RCDC reagents are located in the cabinet of Nancy’s bench.

OXYBLOT PROTEIN
OXIDATION DETECTION

I. Sample Lysis:

Preparation of buffer:

Make up Lysis buffer (store in cold room – can make as a concentrated stock and dilute for use)

· 50mM Tris-HCL (ph8.0)

· 150mM NaCl

· 1% NP-40

The day before:

Freeze water in the cooling chamber of the transfer apparatus
Make up blocking buffer according to chemiflouresence kit instructions
Label tubes for lysis (1 2059 tube if tissue is to be homogenized on polytron + 3 microfuge tubes

Immediately before:

Calculate the amount of lysis buffer needed for experiment and add 100ul of each of phosphatase inhibitor cocktail I + II & 1 complete mini protease inhibitor tablet per 10ml of lysis buffer immediately before use.

Preparation of samples:

Cell culture:

Wash samples 2x in ice cold PBS
Add 1ml/100mm plate of lysis buffer
Scrape samples into microfuge tube

Animal tissues:

Add tissue (fresh or frozen) to lysis buffer in 2059 tube at 150ul minimum/50mg tissue. Immediately homogenize on polytron or equivalent – keeping cold.

All:

Let sit on ice for 30 minutes
Vortex for 30 seconds
Spin in refrigerated centrifuge (cold room) at 10,000xg for 15 min
Transfer 25ul of supernatant to a new tube for running the BCA or RCDC assay to determine protein concentration (refer to the BCA assay or RCDC assay protocol to determine protein concentration).
Transfer the rest of supernatant to a new microfuge tube & store at -80°C
Concentrate (using YM3 column which holds 500ul max) or dilute samples to 2mg/ml for running 10-30ug on a gel

II. Western:

Preparation:

1) cut 3mm filter papers and PVDF membranes (wear powder-free gloves) to the size of the transfer sponges.

2) Obtain enough gels from biochem stores

3) Make up 12% SDS in ddH20

4) Make up 10x gel running buffer

· 30.3g Tris base

· 144g Glycine

· 10g SDS

· Bring to 1 liter with ddH20

5) Make up Transfer buffer (store in cold room)

· 3.36g Tris

· 14.4g Glycine

· bring to 800ml with ddH20

· add 200ml Methanol

Cassette Assembly:

*Using BIO-RAD Mini-PROTEAN 3 Electrophoresis Aparatus.

Remove the Ready-to-go Gel from the storage pouch
Rinse the gel with dH2O and dry it with paper towel
Cut along the dotted line at the bottom of the gel, and pull the clear tape at the bottom to expose the bottom edge of the gel
Refer to the Mini-PROTEAN 3 Electrophoresis Module Assembly guide for assembly instruction

Sample loading:

Calculate the volume needed for 20 ug of protein, and volume for 5x SDS loading buffer
Heat samples at 80°C for 5 mins
Load samples onto gel
Run at 200V for 35-40 mins

Gel Transfer:

Refer to Mini Trans-Blot Electrophoretic Transport cell Instruction Manual
Run at 100V for 1 hr

DNPH Derivatization

*Continuous shaking during all steps

Remove PVDF membrane, and immediately immerse into 100% MeOH for 1 min
Incubate the membrane in 20% MeOH-80% PBS for 5 min
2N HCL for 5 min
Incubate membrane in 2,4-dinitrophenylhydrazine (100ug/ml) in 2N HCL for exactly 5 min
Wash membrane 3x 5 min in 2N HCl
Wash membrane 5x 5 min in 100% MeOH

Immunostaining

Incubate membrane in PBS for 5 min
Block with 5% milk overnight at 4°C
Wash 3x 5 min with 0.05% tween in PBS
Incubate membrane with primary antibody of 1:150 dilution using 5%milk and 1% tween at RT for 1hr
Wash membrane 1x 8 min and 2x 2 min with 0.05% tween in PBS
Incubate membrane with 1:500 secondary antibody using 5%milk and 1% tween at RT for 1hr
Wash membrane 3x 5 min with 0.05% tween in PBS
Place a piece of Saran Wrap on the bench and smooth out, ensure no air bubbles. Pipette 1ml ECF substrate/PVDF membrane onto the SaranWrap.
Lay the blot, protein side down. Incubate for 1 min at RT
Scan the membrane on the Phosphoimager

Section 7 Protocols to evaluate telomere length

Preparation of Tissue for Telomere Probe

Light fixation of fresh or frozen tissue

Please note that fresh or frozen tissue is considered an infectious agent (risk of HIV or hepatitis). Wear gloves, eye protection, and a lab coat when handling tissue. Treat all waste with 10% bleach at least 30 minutes in the bucket in TC marked Biohazard Decontamination

1. Rinse tissue three times in two to three mls of cold PBS.

2. Place tissue in 2 mls of 1% paraformaldehyde for 30 min.

3. Rinse three times in 2 mls PBS.

4. Fix overnight in 70% ethanol at 4° C.

5. Place tissue in LABELED cassettes and take to the appropriate lab to be embedded in paraffin.

a. Don't take tissue to the histochemistry on Fridays. They are too busy to deal with it.

b. If you get tissue on Friday you have two choices. Process it and leave it in 70% ethanol until Monday and then take it to the Histochemistry lab or freeze it in freezing media until it can be processed in a timely manner. The latter option is preferred.

c. Fill out a form and list each cassette. Attach large form with red letters that says: Do NOT Fix in Formalin!! Do NOT Fix in Methacarne!!.

d. Follow the procedure for making sure the tissue does not go into fixative.

e. Blocks are filed in the Nitrogen container near the slide moat moat. Record the location of the block in the log book.

6. When you need sections cut from these blocks, remove them and make a note in the logbook. Take the blocks to the histology lab so that sections can be cut. Make sure one H& E section cut for reference.

7. When you pick up the slides, to return the blocks to the appropriate slots. Put the H & E slides in the H& E box and record in the log book.

Unstained slides are baked at 90°C for 30 min and then deparaffinized as below.

Slide Deparaffinization

1. Place tissue on heat block at 90^oC for 30 minutes.

2. Wash in xylene 3 x 5 minutes at room temperature. Dispose of xylene in waste bottle in hood.

3. Wash in 100%, 90%, 85%, 70% for 3 minutes each at room temperature, with a final brief rinse in water. Air dry and store in liquid nitrogen tank.

Hybridization PNA and Amadate Probes

Ref: Lansdorp, P.M., Verwoerd, N.P., van de Rijke, F.M., Dragowska, V., Little, M.T., Dirks, R.W. Papp A.K., and Tanke, H.J. 1996. Heterogeneity in telomere length of human chromosomes Human Molecular Genetics 5(5):685-91.

Slijepcevic, P., Hande, M.P., Bouffler S.D., Lansdorp, P., and Bryant P.E. 1997. Telomere length, chromatin structure and chromosome fusigenic potential Chromosoma 106(7):413-21.

Before beginning:

Turn on one slide moat to 37°C and a second slide moat to 78°C

Turn on waterbath to 85°C

Make and warm pepsin to 37°C in a Coplin jar in a waterbath

Thaw RNase in ice bucket

Thaw and make working PNA/Amidate solution.

BioRad http://128.95.114.51:8086/confocal/d02/002/2001?display=M&style=B&positioning=S

Lieca https://depts.washington.edu/keck/cgi/calendar/calendar.cgi

Morning:

1. Sections - These will either be embedded in paraffin or frozen sections.

Paraffin embedded lightly fixed – Incubate in 10mM sodium citrate pH 6.5 for 8 min at 85°C. Briefly dip slides in ethanol series (25%, 50 %, then 95%) and air dry.

Frozen sections – Put 50mL of 2% Paraformaldehyde on slide, place cover slip over and let sit for 10min. Rinse with 1X PBS pH 7.2 for 1 min. Briefly dip slide in 25%, 50%, then 95% ethanol. Air-dry completely.

2. Treat with working pepsin solution (heat to 37 °C) for 2 min.

3. Rinse with 1X PBS for 1 min.

4. Dip slides through ethanol series (25%, 50%, then 95% ethanol) and air dry.

5. Add 80 mL of RNase to each slide, cover with coverslip and let sit for 10 min at 37 °C.

6. Rinse with 1X PBS for 1 min.

7. Ethanol series and air-dry.

8. Cover slide with hybrid well. Seal the hybrid well by pressing the edges firmly with the special popcicle stick

9. Add 100 mL of working PNA solution to each slide.

10. Heat slides to 78°C for 10 min.

11. Cover or put slides in dark place, at RT, for 3 hours (overnight is better).

Next day

1. Rinse off probe with 70% Formamide buffer 4 x 15min. It is a good idea to work with the formamide under the fume hood. Waste can be put down the drain but must be recorded on the waste sheet by the drain.

2. Rinse with Tween 20 buffer 4x 5 min.

3. Drain and dip slides through ethanol series and air dry.

4. Add 80 ml DNA counter stain for 5 min (TOTO-3).

5. Wash with PBS for one minute, ethanol series, and air dry.

6. Add 20mL Vectashield (viscous – pipette slowly ) and coverslip

7. If possible, look at with confocal or slides can be stored in –20°C.

Solutions

10 % Blocking reagent (Store at 4 °C in refrigerator 1)

Dissolve 5 grams blocking reagent (cat. # 1096 176 Roche)

In 50 mls of 150 mM Maleic acid, 150 mM NaCl, pH 7.5
(adjusted with NaOH) on heating block. Autoclave and store at 4°C

70% Formamide Buffer for rinsing one liter (Store at RT)

10 mL 1M Tris pH 7.2

290 mL ddH₂O

700 mL Formamide

70% Formamide Buffer with Blocking reagent (Store at RT)

10 mls 10% Roche blocking agent (see above)

90 mls Formamide Buffer

Tween 20 Buffer (Store at RT)

3 L 1X PBS

1.5 mL Tween 20 (pipette very slowly – viscous)

Pepsin (store aliquots at –20°C Freezer 4 Rack 7 )

Stock Pepsin (10 mg/ml)

100 mg pepsin

10 ml dH₂O

1% Working solution

1:100 dilution into .01M HCl

400 mL pepsin stock

39.6 mL .01M HCl

Heat to 37°C

RNase Stock/working 10mg/mL store aliquots at –20°C
In Freezer 4 Rack 7

100 mg RNase

10 mL sterile water

Boil for 10 min to destroy DNase activity

PNA probe FITC and Texas Red both are telomere specific

Stock should be adjusted to 1 ug/ul ~ 100uM

Stock FITC PNA in Refrigerator 2 bottom drawer in a foil wrapped container. We are using this at 1:250 diluted in the formamide blocking solution

Stock Texas Red PNA is in Freezer 4 Rack 4 Box 3

Amidate probe (store aliquots at –20°C) NEEDS Mg++

Amidate stocks are at 500 mg/ml in TE (a 260 OD at 16 = 500 mg/ml)

104 is Centromere probe with FITC label (Freezer 4 Rack 6 Box 3)

104R is Centromere probe with TAMARA label
(Freezer 4 Rack 6 Box 3)

67 is a telomere probe with FITC label (Freezer 4 Rack 6 Box 5)

Working solution:

We are using all three probes at 1:300
300 mL of 70% formamide with blocking solution and 1 mL of probe

5 mL of 500 mM MgSO₄or MgCl₂(You may need to shakethe bottle before using)

PNA/Centromere Probe solution:

Per 3 slides 300 mL 70% fomamide with blocking solution
5 mL 500 mL MgSO₄ or MgCl₂

1 mL PNA probe

1 mL Centromere probe

10 mM Sodium Citrate pH 6.5 (store at RT)

For 1 liter:

2.94 g Sodium citrate

800 mL water

pH to 6.5 with 4N HCl

Bring up to 1 liter with water

.01 M HCl (store at RT )

1 mL of 4M HCl

400 mL of dH₂O

Make 1M HCl : 8.62 mL of concentrated HCl + 100 mL dH₂O

Dilute to .01 M (5 ml 1M HCl : 495 mL dH₂O )

TOTO-3 stock solution is at 1000 mM

Working solution is a 1:200 dilution in PBS

Stock kept in freezer 2

Confocal Microscope set up for photos

General guidelines for Leica

https://depts.washington.edu/keck/cgi/calendar/calendar.cgi

1. Take all pictures at 40x or 100x

2. Format 1024 x 1024

3. Keep a record of the confocal settings used on that day

4. Determine a sub saturating PMT setting for Green (PMT 1) and Red (PMT 2) on bright telomeres and centromeres. Do this with the Glowover –Glowunder setting. Do not change the setting once you have begun to take pictures. If it is obvious the settings you initially choose are not going to work then you must reshoot ALL the slides at those settings

5. Save all images in the Rabinovitch folder in the KECK computer. Then move them ASAP to the U drive.

To move them while you are in the KECK center do the following:
a. Click on the Start icon in the lower left hand corner of the screen.
b. Click on the Run icon in the Start menu.
c. Type the following in the Open space \\fileserver3.pathology.washington.edu\Lab_Rabinovitch and Click OK.
d. You will then be asked for your user and password. For your user name type pathology\your name (the name you use to login on Path computers). For your password, use your current path password. You will then be in the U drive.
f. Then on the screen of the computer in the KECK center use My Computer to find your files.
g. Then move them.

6. To move your files when you are back in the lab do the following:
a. Click on the Start icon in the lower left hand corner of the screen.
b. Click on the Run icon in the Start menu.
c. Type the following in the Open space \\128.95.245.42 and Click OK.
d. You will then be asked for your user and password. Use Rabinovitch and then the lab password
e. You will then be in the KECK computer.
f. Then use My Computer to find the destination you want for your file.
g. Then move them.

7. You are responsible for cataloging your photos so that you can retrieve them..

General Guidelines for BioRad (scanning confocal microscope in the Immunology Cell Analysis Facility)

http://depts.washington.edu/immunweb/cellanalysis/scheduling.html

IP address: 125.95.114.227 (for ftp-ing files to our computer)

** Never turn the BioRad box off while the software is open **

1. Turn key on laser box so the lights are on. (This is usually already turned on for us because it needs to warm up for ~20 minutes prior to use.)

2. Turn on mercury arc lamp.

3. Turn on BioRad box (down is ON).

4. Log in to computer – if this a first-time use, get a username and password from Michelle or Fred.

5. Double-click on LaserSharp2000 Software icon on desktop.

6. Log in to software. (If this is the first time the software is opened during the day, it will take a few minutes (up to 15) to find the laser.) – This may be different from the username and password used to logon to the computer.

7. Check that the joystick is controlling the x-y stage properly. If it is not, close LaserSharp and restart.

8. It should open with the appropriate method for looking at telomere/centromere probe. If not, go to “Methods” and select the appropriate method.

9. Check your light excitation and filter paths.

a. Select the “light path” icon (2^nd from the right) on the controller panel onscreen. This will bring up the light paths. We need to check each individual sequence since we will be scanning sequentially rather than simultaneously.

b. Select “Seq1” – check that PMT 1 is highlighted

i. 568 excitation

ii. B1 – Beamsplitter

iii. 585 EFLP

c. Select “Seq2” – check that PMT 2 is highlighted

i. 488 excitation

ii. B1- Beamsplitter

iii. T2A-560DF

iv. 522 DF 35

d. Select “Seq3” – check that PMT 3 is highlighted

i. 647 excitation

ii. B1 – Beamsplitter

iii. 680 DF 32

10. To look through the oculars, push the filter assembly in the middle of the microscope all the way to the left (blue light should come out through the objective) and push the lever on the bottom left all the way in (to the right) – Find your tissue using the 40x (needs oil) objective. Then switch to the 63x (needs oil) objective.

11. Make sure the “63x” objective and the “1024x1024” format is selected on the controller panel.

12. Test each laser individually to determine saturation and contrast of each color – make sure you are using “x4” speed and “Direct”. Other things to remember:

a. Don’t turn the “Iris” higher than 2.5.

b. Laser power 10 usually gives enough fluorescence. Occasionally turn it up to 30, but risk photobleaching faster.

c. Red == oversaturated; Green == undersaturated (black) – use “offset” to get a balance of these two. Usually don’t want to oversaturate too much.

Do not change the setting once you have begun to take pictures. If it is obvious the settings you initially choose are not going to work then you must reshoot ALL the slides at those settings.
Acquisition: Once the settings are adjusted for the slide, to acquire a picture,

Change speed to “Normal”
Change to “Kalman” and 8 scans – if on the 8^th scan, the image is still changing, increase the number of scans.
Hit the “Sequential acquire” button (2^nd from the left) – each laser will scan through and resolve the pictures based on the number of Kalman scans you make.
The fourth picture is a composite of the three colors you are looking at – RIGHT click the mouse and “Export” à D:Drive à Experiments à Jeannel à 0_Tif_images. Save the file as the number on the slide plus a letter A-J.
You must also save the image as a file; “Save” à number on the slide plus a letter A-J. By hitting save, you will automatically be saving it to your profile folder on the D:Drive.
You must then open a NEW scan.

Protocol For Image Analysis Using
Optimas 6.51

Open Excel and Power Point and minimize
Open Optimas
Go to File click on Open Image File
Open file from U/PUBLIC/confocal images/2004/hyperplastic polyps/# of image
Go to Macro and click on Run
Look in Macros and open june19tissue.mac (You only have to click Run the first time using Optimas in a session, every consecutive time after you can click Run Last. Must click Run again when exit the program and re-open later)
Is there centromere staining: click YES
Enter color code of DNA: click OK
DNA Blue: click OK
Red=1, etc: Click YES
Is DNA segmentation ok: click OK
Click ok to select group 1: click OK
Instructions: click OK
While holding down the “A” button, use the mouse to click all the epithelial cells that are dark blue. They can have pink dots, but must be uniform in color with none of the green or yellow background showing through.
While holding down the “B” button, use the mouse to click on the neighboring stromal cells that surround the chosen epithelial cells. Notice that they are highlighted in yellow instead of white. Again, only choose cells that have a uniform dark blue color, pink dots are okay but no green or yellow background should be showing through.
To un-click either an epithelial or stromal cell just click the cell again and it should un-highlight.
When you are finished choosing cells right click the mouse.
Click ok to select group 1: click CANCEL
Evaluate DNA Zoom: click CANCEL
DNA segmentation ok: click OK
Evaluate Telomere Zoom: click CANCEL
Telomere segmentation ok: click OK
Evaluate Centromere Zoom: click CANCEL
Centromere segmentation ok: click OK
Data is ready: click YES, make sure that Excel IS open before saying yes
Open Excel, go to File and choose Save As
Save in U/PUBLIC/telomere/hyperplasticpolyps and create a new folder for each photo batch
Close the newly saved Excel document and open a blank one and minimize it.
Double Click the shortcut to Irfan View and open Options
Choose Capture
Select Client Area and highlight Optimas in the Menu and click start
Highlight the picture including the title bar and choose copy
Paste the picture on a blank slide in Power Point and adjust the size
Select File and Click Save As
Save all the photos in a single batch in U/PUBLIC/confocalimages/2004/hyperplasticpolyps/Date of Picture/ # of photo batch

EXCEL MACRO INSTRUCTIONS FOR THE ANALYSIS OF OPTIMAS DATA

(protocol updated by Rosana, March 17, 2004)

· Open macro: U:\Public\telomere\colon\MsSummary

· Enable macros? Yes

· Control S. Include just ratio in the summary? Y (appears by default) OK

· Destination file is not present. Choose following: 1 (create new) OK

· Enter the name of the destination file: name with the number and the case – summary

· Browse for the Excel files and select all files in the case, click OPEN

· Choose Method: choose method 1 and click OK

· Choose Threshold: A should be selected already, click OK

· The program should calculate the average for telomeres, centromeres, and ratios of the two, etc. and produce a final Excel sheet

· Copy the final data and paste it into U:\Public\telomere\hyperplastic polyps\hyperplastic polyps final.xls and SAVE

· Close all of the Excel windows for the individual files of the case and save changes

· When closing MsSummary click SAVE ALL

TERMINAL RESTRICTION FRAGMENT (TRF) PROTOCOL

(Rosana Risques, 4/7/04)

DNA EXTRACTION

Extract genomic DNA. The best methods to obtain telomeres in good shape are Phenol-Chloroform extraction or Amersham kit (GenomicPrep Cells and Tissue DNA Isolation Kit #27-5237-01). Freeze-thaw cycles make the telomeres sticky, therefore keep the DNA at 4°C if you have to reuse it in the following days (if not keep it at -20°C). Run DNA in agarose gel to confirm quality. Measure in spectrophotometer or fluorimenter and adjust to 340ng/ul.

DIGESTION

Digest 20ug (10ug x2 tubes) of DNA with either Hinf1 and Alu1, Hinf1 and Msp1 or Hinf1 and Rsa1. (We use Hinf1 and Msp1 from New England Biolabs)

After digestion pool the 2 tubes of each samples and quantitate the genomic DNA in spectrophotometer or fluorimeter (2ul of DNA + 78ul of water). Concentration should be around 200ng/ul (total of 80ul). Run 0.7% agarose gel in 1X TAE to check that the digestions worked. Load 200ng (aprox.1ulDNA + 3ul water + 4ul LB2X).

ELECTROPHORESIS

Make a 0.7% agarose gel in 1XTAE. Load around 2-4ug of DNA (20-40ul). Use P32-labeled markers. Run 16-18 hours at 60v. (enter DNA at 120v. for 3min).

Klenow is from Invitrogen (5u/ul). Dilute to 0.5u/ul in provided dilution buffer.

10X buffer is 10X React2 buffer. lDNA/HindIII is Gibco. 1Kb ladder is Invitrogen.

Purify markers with Qiagen purification kit. Use only 600ul of buffer PE because 700ul tends to spill and contaminates the centrifuge. Elute in 15+15ul of water. Pool the 2 tubes. Take 1ul to read in the scintillation counter. Estimate cpm/ul. Load 100.000cpm in each well.

4. SOUTHERN BLOT

Once the gel has run take a picture of it without taking it out of the box (minimize the risk of breaking it!). Then move it to a tray to do the washes:

- 15 min in depurination sol.

- 30 min in denaturation sol.

- Prepare the blot as shown in the picture. Cut a triangle in the upper left corner in both the membrane and the gel to keep track of the orientation. Don’t forget to put saran wrap around the gel to prevent the liquid to move towards the paper towels. Soak the sponges in the same denaturation sol. used for the gel. Cover the sponges with saran wrap to keep the moisture.

- Blot ON (min: 6 hours, max: all the weekend, as long as it’s well wrapped).

Membrane is Hybond-N from Amersham Biosciences.

After blotting:

- UV crosslink the membrane: 100mJ/cm². Use the UV-crosslinker from Stratagene (Fausto lab) and press ‘autocrosslink’ = 120mJ (membranes usually are around 120cm², so that’s fine). This step immobilizes the DNA in the membrane. If the membrane is charged is not necessary, but it doesn’t hurt.

- 15 min in neutralizing solution

- Put the membrane on Whatman paper to blot the excess of liquid, let it air dry few minutes and cover with saran wrap until pre-hyb.

PREPARATION OF THE PROBE

Probe is a Tel C oligo: CCC TAA CCC TAA CCC TAA CCC TAA (from Qiagen, dissolve in water to make 100pmol/ul and then dilute to 5pmol/ul). To have adequate amounts of probe (i.e. 1-5x10⁶ cpm/ml) label 15pmol of oligo. Run duplicates.

Use gamma-P32-ATP from ICN: >7000 Ci/mmol (2mCi, 120ul).

T4 polynucleotide kinase is from New England Biolabs.

Purify the probe with Microspin G25 colums (from Amersham):

- reaction is 20ul and the recommended volume is 25-30ul, so add 10ul of 10mM Tris-HCl to increase volume up to 30ul

- apply to the columns and spin down (follow manufacturer’s protocol)

- Collect the volume aprox 30ul x2= 60ul.

- Take 1ul to measure in the scintillation counter. The hyb solution is 20ml, so I need 1-5x10⁶ cpm/ml x 20ml= 20-100 x10⁶ cpm.

PRE-HYBRIDIZATION AND HYBRIDIZATION

- Warm pre-hyb and hyb solutions at 37°C

- Cut mesh (#H9088, Apollo, CLP) size of membrane

- Pre-wet in 2X SSC

- Roll in bottle, using 2X SSC to get rid of bubbles

- Discard 2X SSC and add warmed pre-hyb solution

- Incubate in hybridization oven at 37°C for 2h. Spin the bottles in the opposite direction that the membrane was rolled. If not it will roll over itself.

- Denature probe: 5 min in boiling water and put on ice

- Add probe to the warmed hyb solution

- Remove the pre-hyb solution and add hyb solution.

- Incubate in hybridization oven at 37°C overnight (again make the spinning opossite to the direction the membrane was rolled).

WASHES

- Rinse in 5X SSC + 0.1% SDS, RT

- Wash 3X in 0.1X SSC + 0.5% SDS, RT for 8 min.

- Put membrane in Whatman paper to remove excess of liquid. Wrap it in saran wrap.

- Expose in Phosphorimage screen for few hours-days, depending on radioactivity (check signal with Geiger counter).

Q-PCR for Telomeres

Ref. Cawthon RM.  Telomere measurement by quantitative PCR,  Nucleic Acids Res. 2002 May 15;30(10):e47

Ref. Cawthon RM. Conditions for quantitative PCR of telomere length, August 7, 2003.

This protocol is used to determine relative telomere length. In order to do this you must run each set of samples with a different set of primers. The first set is 36B4 primers for acidic ribosomal phosphoproteinPO 1. This is a single copy gene. The second set of primers, tel 1 and tel 2 amplify the telomere length. The relative telomere length is determined by a ratio of the PCR product of the single copy gene and the PCR product of the telomere.

The primer sequences written 5'3':

tel 1b, CGGTTTGTTTGGGTTTGGGTTTGGGTTTGGGTTTGGGTT

tel 2b, GGCTTGCCTTACCCTTACCCTTACCCTTACCCTTACCCT

36B4u, CAGCAAGTGGGAAGGTGTAATCC

36B4d, CCCATTCTATCATCAACGGGTACAA

36B4 = acidic ribosomal phosphoproteinPO located on chromosome 12

Q-PCR (for telomere PCR)
		MIX
	1X	100X	dilution	final conc.	Primers:
6-ROX 50X	0.40	40.00	1/50	1X	tel 1b	300nM
SybrGreen I 10X	0.80	80.00	1/25	0.4X	tel 2b	300nM
Buffer 10X	2.00	200.00	1/10	20mM Tris-HCl pH 8.0 + 50mM KCl
MgCl₂ 50mM	0.60	60.00	1/33.3	1.5mM
dNTPs 10mM	0.40	40.00	1/50	0.2mM
DTT 100mM	0.50	50.00	1/40	2.5mM
DMSO 100%	0.20	20.00	1/100	1%
Taq (5u/ml)	0.16	16.00		0.8u
Tel 1b 3uM	2.00	200.00	1/10	300nM
Tel 2b 3uM	2.00	200.00	1/10	300nM
H₂O	8.94	894.00
DNA (ng/ml)	2.00			adjusted
Check Sum	20.00	1800.00
		18.00

Q-PCR (for single copy gene PCR)
		MIX
	1X	100X	dilution	final conc.	Primers:
6-ROX 50X	0.40	40.00	1/50	1X	36B4u	300nM
SybrGreen I 10X	0.80	80.00	1/25	0.4X	36B4d	500nM
Buffer 10X	2.00	200.00	1/10	20mM Tris-HCl pH 8.0 + 50mM KCl
MgCl₂ 50mM	1.40	140.00	1/14.29	3.5mM
dNTPs 10mM	0.40	40.00	1/50	0.2mM
DTT 100mM	0.50	50.00	1/40	2.5mM
DMSO 100%	0.20	20.00	1/100	1%
Taq (5u/ml)	0.10	10.00		0.5u
Primer a 10X	2.00	200.00	1/10	300nM
Primer b 10X	2.00	200.00	1/10	500nM
H₂O	8.20	820.00
DNA (1ng/ml)	2.00			adjusted
Check Sum	20.00	1800.00
		18.00

Notes:

1) The taq being used is Platinum TaqPCRx DNA Polymerase purchased from Invitrogen (Cat. # 11509-015).

2) 6-ROX is purchased from Invitrogen (Cat. # 12223-012).

3) SybrGreen is purchased from Molecular Probes (Cat. # S-7563).

4) Amount of mix can be adjusted depending on number of samples.

5) Amount of DNA can be adjusted depending on experiment.

Methods of Analysis:

DDCt Method

A reference sample is used as the calibrator. The following set of equations are used in the determination of 2DDCt:

DCt = Ct(tel) - Ct (36B4)

DDCt = DCt(calibrator) - DCt(sample)

2DDCt = 2^DDCt

The magnitude of 2DDCt is a measure of the telomere length of the sample. A larger 2DDCt implies a longer telomere.

Example of Plate Setup

1	2	3	4	5	6	7	8	9	10	11	12
1	2	3	4	5	6	7	8	9	10	11	12
1	2	3	4	5	6	7	8	9	10	11	12
13	14	15	16	17	18	19	20	21	22	23	24
13	14	15	16	17	18	19	20	21	22	23	24
13	14	15	16	17	18	19	20	21	22	23	24
Ctr.1	Ctr.1	Ctr.1	Ctr.2	Ctr.2	Ctr.2	Ctr.3	Ctr.3	Ctr.3	NTC	NTC	NTC
X	X	X	X	X	X	X	X	X	X	X	X

The setup of the plate can be adjusted according to the experiment. The unknowns should at least be in triplicates to ensure accuracy.

Numerical numbers (1-24) represent the unknown DNA samples. Ctr.1-3 are 3 different DNA controls of known relative telomere length respectively. By Q-PCR, the telomere lengths of the unknown DNA samples can be estimated by comparison with that of the DNA controls.

PCR protocols:

For tel primers			For 36B4 primers
Temp	Time	Cycles	Temp	Time	Cycles
95°C	10 min	one	95°C	10 min	one
95°C	15 sec	30	95°C	15 sec	40
56°C	1 min	cycles	56°C	20 sec	cycles
			72°C	20 sec

Remember to add dissociation stage.

Last updated 04/30/2004, KL

Using ABI PRISM 7900

Set-up

Open SDS 2.1 Program (double click on SDS 2.1 icon on desktop)
Click on File à New
Under Container, select 96 Wells Clear Plate, then click OK
Select all wells by clicking on the square at the top left hand corner of the picture of the plate
Click on Tools à Detector Manager
Select the detector used, click on Copy to Plate Document, then click on Done
Check the box under Use next to the selected detector
Click on the Instrument tab
Enter Sample Volume
The box next to 9600 Emulation should be checked (this should be the default)
Click Add Dissociation Stage
Enter time, temperature, number of cycles in the Thermal Profile
Make sure Ramp Rate of the last step of the dissociation stage is 2%, while other steps are 100% (this should be the default)
Make sure the date collection symbol shows up in stage 1, stage 2, the last ramp and last step in the dissociation stage (this should be the default)
Save file by clicking File à Save As
Click Open/Close button
Put sample plate in the machine. Remember to have pad on top of plate to prevent evaporation
Click Open/Close button again
Click Start button

Analysis

Select all wells by clicking on the square at the top left hand corner of the picture of the plate
Click on Tools à Detector Manager (skip steps 21-26 if detector already has the correct name)
Click on New
Type in Name, select the right Reporter and choose Quencher to be Non Fluorescent. Click OK. This is just to rename the detector
Select this new detector, click on Copy to Plate Document then click on Done
Uncheck the box under Use next to the original detector
Check the box under Use next to the new detector
Select the wells for non-template control. Click on Task cell next to selected detector and choose NTC
Select the wells for each replicate in the standard curve. Click on Task cell next to selected detector and choose Standard. Enter amount under Quantity
Do the same for the other replicates in the standard curve
Select the other wells. Click on Task cell next to selected detector and choose Unknown
Click on Analyze button in the Toolbar or click Analysis à Analyze
Adjust Baseline and Threshold so that R^2 in the standard curve is as high as possible and the slope of the standard curve is near –3.3
To export data, click on File à Export. Export as a .xls file for analysis in Excel.

Last updated by KL 04/30/04.

Section 8 BAC cloning protocols

BAC Prep

(Bryce Sopher, La Spada lab, Clontech Nucleobond AX-500)

1. Inoculate 3 ml overnight culture from single colony on plate.

2. Dilute 1 ml of overnight culture into 250 ml culture.

3. Grow to A₆₀₀ of 1.0.

4. Pellet bacteria by centrifuging at 5K, 10 minutes. Continue, or store pellet frozen at –80C.

5. Carefully resuspend the bacterial cell pellet in 12 ml buffer S1 + RNAse.

6. Add 12 ml buffer S2. Mix gently by inverting tube and incubate at room temperature for 5 minutes. Do not vortex.

7. Add 12 ml buffer S3. Mix gently by inverting 6-8 times until homogenous suspension is formed. Incubate on ice for 10 minutes to precipitate SDS and cellular debris.

8. Meanwhile, equilibrate Nucleobond AX-500 cartridge with 5 ml buffer N2.

9. Put folded filter into funnel over equilibrated AX-500 cartridge and wet with ~1 ml H2O.

10. Pour ice-cold suspension into filter/funnel and load onto equilibrated cartridge. Collect flow-through in 50 ml tubes (waste).

11. Wash cartridge 2X with 12 ml buffer N3.

12. Elute DNA into 15 ml tube with 7.4 ml buffer N5.

13. Precipitate DNA with 5.1 ml (0.7 vol) of room temperature isopropanol.

14. For increased yield precipitate ON at 4C.

15. Centrifuge for at least 1 hr at max speed at 4C in table top swinging bucket rotor.

16. Wash pellet with 70% EtOH and centrifuge at least 1 hr at max speed at 4C.

17. Pour off EtOH and air dry pellet briefly.

18. Resuspend pellet in 100 ml microinjection buffer.

Plasmid Mini-Prep:
Preparation of BAC DNA for analytical purposes.

Pick colonies and inoculate 2ml of LB culture containing the appropriate antibiotics. Incubate at 37˚C overnight with shaking at 1100rpm.

Next Day –

Transfer overnight culture to a 1.5ml to eppendorf tube.

Spin down the tube for 2 min. at 11,000 rpm.

Discard the supernatant and resuspend the cell pellet in 200 ml buffer P1 with RNase (from Qiagen DNA Maxi-preparation kit). Gently mix the buffer and break up the pellet without making too frothy.

Add 200ml of buffer P2 (Qiagen) and quickly mix by inverting the tube several times (Count to 6). Incubate at RT for 5 mins. Solution will become viscous and clear.

Add 200ml of buffer P3 (Qiagen) and quickly mix by inverting the tube several times. Solution will become gloppy and have big white globs. Incubate on ice for 5 mins.

Spin down the white lysate at the highest speed for 5 mins.

Transfer the clear supernatant into a new 1.5ml eppendorf tube and add 0.60ml of 2-propanol (iso-propanol).

Mix by inverting the tube. PPT in fridge for 1hr. Spin down the DNA at highest speed for 10 mins.

Discard the supernatant and add 1ml of 80% ethanol to rinse the pellet. Be careful not to loose the small white pellet. Don’t even loosen it (If you do loosen it, spin down for another 1-2 mins.).

Clean the inner wall of the tube with a piece of tissue or cotton stick.

Dry the pellet under the speed vacuum for 2 mins. Or leave the tube open on the bench for 5 to 10 min until the DNA pellet is completely dried. DO not over dry to pellet otherwise the DNA will become difficult to re-dissolve.

Resuspend pellet in 20ml of TE.

Spec on spectrophotometer – 1:200 dilution. Store in –20˚C chest freezer.

Transformation

– to prepare cells to shuttle/carry your plasmid of interest

1. To a 2059 tube, add 50ml of competent cells (bought from Biochem stores, stored in –80˚C).

2. Add ~20ng of DNA plasmid.

3. Incubate on ice for 30 mins.

4. Heat shock in 42˚C water bath for 45 seconds.

5. Throw on ice for 2 mins.

6. Add 450ml of SOC medium – SOB media +

7. Incubate at 37˚C for 1 hr. or less.

8. Plate out on LB-AMP plates, grow at 37˚C overnight.

BACs containing GAPDH

62KB GAPDH 43 KB
CTD2224PD (104 KB)

in pBeloBACII from Research Genetics/Invitrogen

http://www.genome.clemson.edu/groups/bac/protocols/protocols2new.html

The complete sequence for pBeloBACII is on the U Drive U:\PUBLIC\BAC Constructs\BAC CTD2224P2\ CTD2224P2 sequence.doc.

Noel Hudson has removed the LoxP site from the CTD2224PD and replaced it with amp^r, The amp^r CTD2224PD will be the target for the mCAT, GCLM, and SOD constructs. The DNA for the amp^r CTD2224PD is in the refrigerator in a 15 ml tube. Jeanne knows where this is. The BAC amp^r CTD2224PD is in DY380 cells on plates in the refrigerator.

It may be necessary to grow this up (30ºC) and do a BAC prep. Digest it and do a pulse field to check to see if the Lox is gone and the amp is there.

We have a second BAC RP11-369N23 which has not been used.

70KB GAPDH 139 KB

RP11-369N23 (210 KB)

in pBACe3.6 from BACPAC Resources

http://bacpac.chori.org/pbace36.htm

The complete sequence for pBACe3.6 is on the U Drive U:\PUBLIC\BAC Constructs\BAC RP11369N23\ RP11369N23 sequence.doc.

The sequence for the insert is

U:\Lab_Rabinovitch\PUBLIC\BAC Constructs\loxP.STOP.loxP.doc (loxPSTOP lox P)

U:\PUBLIC\BAC Constructs\MCAT\MCAT cassette.doc (OCT and MCAT and a portion of the poly A tail)

\\Pathology\Files\Lab_Rabinovitch\PUBLIC\BAC Constructs\pBluescript II SK-.doc (Blue Scipt II from which the pY (p Martini) comes)

The multiple cloning site for pY is in Noel’s BAC notebook page 76.

Section 9 Flow Protocols for Cell Cycle, Proliferation, Viability and Coast Sea Food.

Viable Hoescht 33342 Stain

1. Cells should be maintained in the media that is normal for their growth.

2. Add 10 ml of 1 mM Hoechst 33342/ml of media. Incubate for 30 minutes at 37°C.

3. Cells must be maintained in Hoechst, DO NOT wash out the stain.

4. Run samples on the flow cytometer as soon as possible (with in minutes) of staining.

Hoechst 33342 (viable DNA stain) 1mM

200 ml:

0.112 g Hoechst 33342*

200 ml of dH₂0

2 ml 95% ethanol

          Mix thoroughly. Cover the bottle with aluminum foil to protect from
          the light.
          Store at 4°C.

* CALBIOCHEM cat.# 382065-Q

Bisbenzimide H 33342 Fluorochrome, Trihydrochloride.

BrdU/Ho/Eb

Ref. Rabinovitch, P.S., Kubbies, M., Chen, Y.C., Schindler, D., and Hoehn, H., BrdU-Hoechst Flow Cytometry: A Unique Tool for Quantitative Cell Cycle Analysis. Exp. Cell Res. 174:309-318. 1988.

1. Plate cells at 200,000/25 cm²flask in 4 ml MEM, 0.1% FCS. Old cells should be plated at about 100,000 cells/flask.

2. Purge incubator several times to equilibrate media quickly.

3. Leave in 0.1% FCS for 5 days.

4. On day 5, feed with 3-4 ml MEM media containing 1.5 x 10^-4BrdU M and desired concentrations of FCS (0.1-16% for experimental samples; 10% for tests). You may include deoxycytidine at this step.

5. Feed cells with fresh media every 2-3 days throughout experiment under a sodium vapor light.

6. Sample at various time points (16 hours - 14 days for experimental samples; 7 days for tests).

7. Trypsinize and pellet cells. Resuspend pellet in 1 ml MEM with 10% FCS, 10% DMSO and freeze (not preferred) or resuspend in 0.8 ml Hoechst buffer, can put on ice for at least 2 hours and analyze. Samples that have been on ice for at least 2 hours may also be frozen (add 0.1 ml DMSO while vortexing) and analyzed at a later time. (Samples may also be stained with NEB-FMI instead of Hoechst)

8. After Hoechst stain add stock ethidium bromide for a final concentration of 2.0 mg/ml.

9. Add stock Mg (500 mM MgCl₂) for a final concentration of 5 mM.

Stock 10^-2M BrdU Solution: 0.0921 g BrdU (MW 307.4)

(keep stock for up to 1 month) 30 ml MEM – Filter sterilize before use

Hoechst Buffer: for 500 ml

Final concentration Amount needed for 500 mls

0.154 M NaCl 4.5 g NaCl

0.1 M Tris, pH 7.4 50 ml of 1 M Tris pH 7.4

0.1% NONIDENT P-40 500 ml of NONIDENT P-40

(IGEPAL CA-60)

1 mM CaCl₂ 1 ml of 500 mM CaCl₂

5 mM MgCl_{2 5
ml of 500 mM MgCl2}

0.2% BSA 1 g BSA

2.5mg/ml Hoechst 33258 2 ml of 1.25 mg/ml Hoechst 33358

qs to 500 ml with dH₂0

Stock 10^-2M Deoxycytidine Solution:

(use 0.2 ml/100ml media for 0.0227 g deoxycytidine (MW 227)

final concentration of 20 mM) 10 ml MEM – Filter sterilize before use

See Proliferative Survival Protocol – “Processing data for the Proliferative Survival Protocol” for information about gating and calculation to determine the number of cells in each component of the cell cycle.

BrdU

(Reid/Rabinovitch)

This protocol works well for measuring cells in S-phase.

1. Thaw samples to room temperature.

2. Spin cells to 1500 rpms for 10 minutes, decant.

3. Resuspend in 400 ml 0.1% NP40/PBS + 5% FCS and mince if using tissue.

4. Syringe using a 23G needle; slowly 3 times.

5. Denature DNA.

a. Best results used 4N HCl at 23°C for 40 minutes.

b. Can also use 2N HCl at 37°C for 40 minutes (4 ml).

NOTE: Vortex twice during 40 minutes.

6. Filter through millipore with 70 mm screen. This step is not necessary if you are using cells rather than tissue.

7. At 40 minutes, add 8 ml of Tris butter + 0.5% BSA + 5mM HEPES.

8. Spin at 1500 rpms for 10 minutes, decant.

9. Add 5 ml Tris + 0.5% BSA + 5mM HEPES, vortex.

10. Spin at 1500 rpms for 10 minutes, decant.

11. Add 20 ul BrdU conjugate and 20 ml PBS/5% FCS. Total volume = 50 ml.

12. Incubate 30 minutes at room temperature.

13. Add 5 ml PBS/5% FCS, vortex.

14. Spin at 1500 rpms for 10 minutes, decant.

15. Resuspend 500 ul PBS/5% FCS.

16. Add 5 ml Propidium Iodide stock = 10 ug/ml final concentration.

17. Add 50 ml RNAase 1% stock solution.

18. Incubate 20-30 minutes, RT.

19. Syringe with 25G needle; 3 times.

20. Run on flow cytometer.

Source BrdU conjugate

BrdU - Ab

Single Step BrdU Protocol

1. Spin down cells and decant.

2. Add 1 ml PBS + 5% NGS, spin down cells and decant.

3. QS pellets to 150 ml with PBS + 5% NGS.

4. Add 450 ml Membrane Shredding Solution.

5. Put samples on ice for 15 minutes, vortex periodically (every 3 minutes).

6. Add 600 ml 4 N HCl, agitate for 30 minutes at RT.

7. Add 1 ml of 1 M Tris (pH 7.2).

8. Spin down, resuspend with 3 ml of 1 M Tris (pH 7.2). Spin down, decant.

9. Resuspend in 100 ml of 1:20 FITC conjugated anti-BrdU antibody (from Caltag).

10. Add 1 ml of PBS + 5% NGS, spin down cells and decant.

11. Resuspend in 250 ml of PBS with 10 mg/ml DAPI.

Two Step BrdU Protocol

Adapted from Rissa Sanchez’s protocol

1. Spin down cells and decant.

2. Add 1 ml PBS + 5% NGS, spin down cells and decant.

3. QS pellets to 150 ml PBS + 5% NGS.

4. Add 300 ml Membrane Shredding Solution.

5. Put samples on ice for 15 minutes, vortex periodically (every 5 minutes).

6. Add 75 ml 2 N HCl, agitate for 30 minutes at RT.

7. Add 1 ml of 1 M Tris (pH 7.2).

8. Spin down, resuspend with 1 ml of 1 M Tris (pH 7.2). Spin down, decant.

9. Resuspend in 750 ml 1 M Tris (pH 7.2).

10. Add 20 ml of anti-BrdU primary from DAKO. Keep on ice, 15 minutes.

11. Add 1 ml of PBS + 5% NGS, spin down cells and decant.

12. QS to 200 ml with PBS + 5% NGS.

13. Add 10 ml GAM-PE (Biomeda) resuspended in PBS + 5% NGS.

14. Incubate on ice for 45-60 minutes.

15. Add 1 ml of PBS + 5% NGS, spin down cells and decant.

16. Resuspend in 200 ml of PBS with 10 mg/ml DAPI.

Three Step BrdU Protocol

Adapted from Rob Pierce’s protocol

1. Spin down cells and decant.

2. Add 1 ml PBS + 5% NGS, spin down cells and decant.

3. QS pellets to 150 ml with PBS |5% NGS

4. Add 300 ml of Membrane Shredding Solution.

5. Put samples on ice for 15 minutes, vortex periodically (every 5 minutes).

6. Add 75 ml 2 N HCl, agitate for 30 minutes at RT.

7. Add 1 ml of 1 M Tris (pH 7.2).

8. Spin down, resuspend with 1 ml of 1 M Tris (pH 7.2). Spin down, decant.

9. Resuspend in 500 ml of PBS + 5% NGS containing 10 ml of anti-BrdU antibody. Keep in dark 30’ @ RT.

10. Add 1 ml of PBS + 5% NGS. Spin down cells and decant.

11. Add 25 ml 1:50 secondary biotinylated goat-anti-mouse antibody (1:100 final dilution in 50 ml volume).

12. Incubate in dark for 30’ @ RT.

13. Add 1 ml of PBS + 5% NGS. Spin down cells and decant.

14. Add 25 ml 1:100 strepavidin-PE

15. Incubate at RT for 20’.

16. Add 1 ml of PBS + 5% NGS. Spin down cells and decant.

17. Resuspend in 200 ml of PBS with 10 mg/ml DAPI

Membrane Shredding Solution:
100 ml PBS

500 ml NP-40

20 mg EDTA, pH 7.2

Cytokeratin/DAPI

1. Have ready two eppendorf tubes (5 x 10⁵ cells per tube) for each cell line to be tested. Cells may be suspended in media or PBS. Volume of cells in each tube should be 100 ml or less. One tube will be for a negative control to check for non-specific binding of Ig isotype antibody. For positive control, include a known epithelial cell line (e.g. HeLa or SW-48) in the assay.

2. Treat cells with 0.5 ml of 0.5% buffered formalin solution (dilute 2% stock) for 10 minutes on ice. [Use EM grade methanol-free formalin.]

3. Qs to 2 ml with PBA (1X PBS + 0.1% BSA fraction V). Keep PBA sterile by aliquoting into 15 ml tube.

4. Centrifuge for 8-10 minutes at 200 G at 4°C. Remove supernatant.

5. Add 1 ml of 0.1% Triton PBA (dilute 0.5% stock). Incubate on ice for 3 mins.

6. Qs to 2 ml with PBA and spin. Remove supernatant.

7. For each pair of tubes from each cell line, label one (+) for cytokeratin and one(-) for isotype. Add 10 ml of Anti-cytokeratin FITC (CAM 5.2) (Becton-Dickinson) straight from bottle into (+) tube. For (-) tube (negative control), add 10 ml of Mouse IgG_2a FITC (Becton-Dickinson) straight from bottle. Incubate 45 mins on ice in dark.

8. Qs to 2 ml with 0.1% Triton PBA and spin. Remove supernatant.

9. Add 0.3 ml DAPI.

**All reagents are stored at 4°C

DAPI Staining

DAPI is used to stain DNA and in our group is normally used to determine cell cycle information.

1. In most cases the cells will be spun down and the supernatant removed before adding DAPI.

2. Resuspend the pellet in at least 200 ml of DAPI. In general you would want the
cells at a concentration of 2 x 10⁶/ ml of DAPI. The cells can be run on the flow cytometer immediately.

3. The cells can also be frozen for analysis at a later date.

Fresh cells in suspension:

DNA Content and Cell Cycle Analysis.

Analysis is performed as previously described (Rabinovitch, P.S., Reid, B.J., Haggitt, R.C., Norwood, T.H., and Rubin, C.E. Progression to cancer in Barrett's esophagus is associated with genomic instability. Lab. Invest. 60:65‑71,1989). In brief, cells in suspension are spun and resuspended in a solution of 10mg/ml 4,6-diamidino-2-phenylindole (DAPI) and 0.1% nonidet P-40 detergent in a Tris buffered saline. The suspension is triturated with a 26 gauge needle and analyzed using a Coulter ELITE cytometer (Coulter Corp., Miami FL), with ultraviolet excitation and DAPI emission collected at >450nm. DNA content and cell cycle are analyzed as previously described (Rabinovitch, P.S. DNA content histogram and cell cycle analysis. Meth. Cell Biol. 41:263-296,1994) using the software program MultiCycle (Phoenix Flow Systems, San Diego, CA).

Tissues:

DNA Content and Cell Cycle Analysis.

Analysis is performed as previously described (Rabinovitch, P.S., Reid, B.J., Haggitt, R.C., Norwood, T.H., and Rubin, C.E. Progression to cancer in Barrett's esophagus is associated with genomic instability. Lab. Invest. 60:65‑71,1989). In brief, tissue is minced with scalpels in a solution of 10mg/ml 4,6-diamidino-2-phenylindole (DAPI) and 0.1% nonidet P-40 detergent in a Tris buffered saline. The supernatant is triturated with a 26 gauge needle, filtered through 40mm steel mesh, and analyzed using a Coulter ELITE cytometer (Coulter Corp., Miami FL), with ultraviolet excitation and DAPI emission collected at >450nm. DNA content and cell cycle are analyzed as previously described (Rabinovitch, P.S. DNA content histogram and cell cycle analysis. Meth. Cell Biol. 41:263-296,1994) using the software program MultiCycle (Phoenix Flow Systems, San Diego, CA).

Cells from Paraffin:

DNA Content and Cell Cycle Analysis.

50mm sections cut from paraffin-embedded tissue are processed by a variation of the technique originally reported by Hedley et. al. (Hedley DW, Friedlander ML, Taylor IW, Rugg CA, Musgrove EA. Method for analysis of cellular DNA content of paraffin-embedded pathological material using flow cytometry. J. Histochem. Cytochem. 31:1333-1335, 1983). In brief, sections are dewaxed in xylene, gradually rehydrated in a step series of ethanol solutions, and digested in water with 1% pepsin pH 1.5 37ºC for 40 min. The supernatant is triturated with a 26 gauge needle and resuspended in an isotonic pH 7.4 buffered solution with 0.1% nonidet P-40 detergent, 10 mg/ml diamidino-2-phenylindole (DAPI) and 1% RNAse, and filtered through 40mm steel mesh. The analysis is performed on an ELITE cytometer (Coulter Corp., Miami FL) using UV excitation. 50, 000 cells are analyzed, if available, and in all cases acceptable histograms contained at least 10,000 cells and a coefficient of variation below 6.0%.DNA content and cell cycle are analyzed as previously described (Rabinovitch, P.S. DNA content histogram and cell cycle analysis. Meth. Cell Biol. 41:263-296,1994) using the software program MultiCycle (Phoenix Flow Systems, San Diego, CA).

1X DAPI (Working Solution)

To 500 ml dd H₂O add:

8.5g NaCl (final conc. = 146 mM)

1.2 g Tris Base (final conc. = 10 mM)

Adjust pH to 7.4 with HCl.

Add:

4 ml of 500 mM CaCl₂solution (final conc. = 2 mM)

44 ml of 500 mM MgCl₂solution (final conc. = 22 mM)

50 mg (0.05g) BSA

1 ml nonident P-40 (NOT the same as NP-40) detergent (final conc. = 0.1%)

10 mg DAPI (4,6-diamidino-2-phenylindole*) powder

(final conc. = 10 ug/ml)

100 ml DMSO (final conc. = 10%)

Add dd H₂0 to final volume of 1 L.

Store in dark or foil wrapped bottle at 2-6^oC.

*Accurate Chem.Co. #18860 (no substitutes)

KI-67 staining

Ki67/DNA content Multiparameter Flow Assay for Cell Cycle Proliferation and Cell Sorting (Written by Rissa Sanchez)

I. Materials

A. Ice in ice bucket

B. Underpad, Versidry, (Fisher 1420638)

C. Splash shield

D. Gloves, powder free

E. Microfuge rack

F. Tube, 15 ml., conical (Fisher 05-53859B)

G. Tubes, microfuge, siliconized, RNase- and DNase-free, 1.7 ml, (National Brand through Fisher 11-842-54)

H. Syringes, 1 cc tuberculin with 25 gauge needle (Fisher 14-826-88)

I. Syringe, 20 cc (Fisher 14-829-21B)

J. Filter, 0.22 micron (Fisher DDA0202550)

K. Pipettors, Single channel: 1000 ml, 200 ml, 20 ml

L. Pipette tips, aerosol, ART brand

M. Petri dishes, 35 x 10 mm, bottoms only (Fisher 08-757-100A)

N. Scalpel handles (#9) and blades (#10, Fisher 08-916-5A)

O. Hemostat

P. Serum, normal goat (Caltag #10000)

Q. Ki67-PE antibody (DAKO R0840)

R. IgG1-PE antibody (DAKO X0928)

S. Pipettes, 5ml., transfer, individual wrap (Fisher 13-675-22)

II. Reagents

A. NST buffer (146 mM NaCl, 10mM Tris Base (pH 7.5), 1 mM CaCl₂, 0.5 mM MgSO₄,0.05% Bovine serum albumin (BSA), 21 mM MgCl₂, 0.2% Nonidet P40

B. 100X DAPI (4,6-diamindine-2-phenylindole dihydrochloride) 1 mg/ml (Roche 236276)

III. Safety

Use universal precautions for human tissue including lab coats, gloves, splash shield. Never recap syringes. Discard syringes into biohazard safety containers. Use disposable scalpels with handles and discard in red biohazard sharps containers when finished.

IV. Protocol

A. Control

Lymphocytes stimulated with PHA for 48 hours or a lymphoblast cell line is used as a control for the Ki67 antibody and sorting protocol . Typically, several million cells are grown as a lot and approximately 5 x 10⁵ cells subaliquotted into microcentrifuge tubes and frozen with 10% DMSO. Use one aliquot per experiment. To prepare control cells thaw aliquot, centrifuge at 2000 rpm (200g) on the TOMY centrifuge at 0°C for 10 minutes. Decant. Add 200µl NST to pellet and syringe with a 1cc syringe with 25-gauge needle to enucleate the cells about 5 times. Adjust the volume to 360µl with NST and add 40µl normal goat serum. Divide the cell suspension into two microfuge tubes with a final volume of 200µl each. Tubes are typically pre-labelled as (1), the positive Ki67-PE control, and (2), the negative IgG1-PE control. Keep tubes on ice until mincing of biopsies is complete. This step can be performed either before or after tissue mincing. Use control cells to recheck titer and antibody concentrations between different lots of antibodies.

B. Documentation and set-up

Write flow experiment in lab notebook including patient name, flow number, sample number and level. Note relevant details on the experiment worksheet if different from the norm eg. biopsies that are especially small or necrotic, if the media color is off. Also note the lot number of a new vial of antibody when opened. Label tubes with sequential numbers. Tube 1 is positive lymphocyte control, Tube 2 is negative lymphocyte control. Each biopsy for the day will be assigned three tubes labeled sequentially 3, 4 and 5; and the next patient sample will be assigned 6, 7 and 8, etc. The first of the three tubes is designated for DNA content analysis, the second tube is for the negative antibody and the third tube is for the positive antibody.

C. Biopsy Mincing-Always change scalpel blades with each biopsy.

In the bottom of a petri dish mince biopsy in 600 µl NST buffer. There is an “art” to mincing that requires personal instruction to ensure that the highest yield of nuclei is recovered per biopsy. The mechanics of tissue dissaggregation and enucleation include mincing with scalpels into tissue fragments/suspension that can be aspirated by a 1000µl pipetter. It is important that the pipetter be set at least at 600µl as any volume less that 600µl will not create enough force to shear adequately. The first step is to shear with the 1000µl pipetter several times ~ 7 times. It is very important in all shearing steps to avoid creating aerosols or “bubbles” which will severely compromise total yields. The second shearing step uses a 1cc syringe with a 25-gauge needle. This step is especially vexing as tissue fragments are prone to clog. This step is also one of the most hazardous and requires extreme caution because one is using potentially infectious tissue with a sharp under extreme pressures. NEVER force the suspension through the bore of the syringe if there is a clog in the needle because under pressure the needle will shoot off the end of the syringe spraying suspension. It is best to avoid tissue fragments. If a fragment is collected on the bore of the needle flick it off. Run the suspension through the syringe about 7 times. Transfer the suspension into the patient’s negative control tube. Add another 600µl of NST to the petri dish and shear the suspension as before with both the 1000µl pipetter and 1cc syringe and add to the same tube.

D. DNA Content

For each patient sample, remove 100µl into the tube designated for DNA content. This is important because the antibody staining protocol requires centrifugation that creates aggregates. Given the importance of detecting 4N abnormalities in Barrett’s esophagus it is imperative that nuclei suspensions for DNA content must NEVER be centrifuged. Hold on ice while preparing the other biopsies. The tube with the 100µl suspension of nuclei for DNA content will require another 100µl of NST with DAPI at 10µg/ml final concentration.

E. Antibody staining

Centrifuge all the minced samples at 2000 rpm (200g) on the TOMY centrifuge at 0°C for 10 minutes. Remove proteolytic enzymes that degrade protein by decanting. Decant by inverting the tube in one fluid motion onto the Versiday pad. NEVER tap the tube in an attempt to remove all the fluid because you may loose the pellet. There will always be about 50µl remaining in the tube held by surface tension. The following is a fairly labor intensive distribution of each patient’s sample with the goal of optimizing sorting yield by enriching the number of nuclei into the Ki67-PE tube. Set the 200µl pipetter to 100µl. Add 200µl of NST + 10%NGS to the pellet in the IgG1 negative antibody tube and resuspend with the pipetter. Then transfer 200µl of the nuclear suspension into the Ki67-PE tube for that patient. This tube will have 80-90% of the total number of nuclei harvested from the biopsy. There should about 20-50µl left in the IgG1 tube which typically should satisfy the 15,000 events required for the analysis of the negative sample. Add enough NST + 10%NGS to the IgG1 tube to a final volume of 200µl. After all the samples have been distributed into their appropriate DNA content, negative and positive tubes, appropriate antibody can be added. Currently we add 10µl of the Ki67-PE and 7.5µl of the IgG1-PE to the appropriate 200µl sample. After adding antibody, briefly mix each tube with vortex. Place on ice. Incubate on ice for a minimum of 30 minutes, centrifuge at 2000 rpm on the TOMY centrifuge at 0°C for 10 minutes. Decant as before. Resuspend pellets with NST + 10% FCS + DAPI at 10µg/ml final concentration. Tubes are ready for analysis and sorting.

F. Flow cytometer set-up (see Ki67/DNA protocol stored on Coulter Elite)

G. Analysis and sorting

Immediately before each sample is run on the flow cytometer each sample is syringed again with a 1cc syringe and a 25 gauge needle x 7 and transferred into a 5 ml. labeled falcon tube through a custom made 70 micron prefilter. These prefilters are reused after each use. Prefilters are cleaned with bleach, water washs, alcohol sterilization and air-drying. Prefilters are also periodically checked with a 10x lens and discarded if corroded or damaged. Be sure that these filters are isolated from fibers that may contaminate the nuclei suspension and become trapped in the flow cell! The flow cytometer also has similar custom made prefilters that are changed between patients.

Sorting strategy: If the biopsy is diploid, sort Ki67 positive G1 and all the cells in the 4N fraction. If the biopsy has an aneuploid and a non-cycling diploid, sort all the 2N cells and all the aneuploid G0/G1 cells. If the biopsy has an aneuploid and a cycling diploid fraction, sort the cycling diploid G1 fraction and all the aneuploid G0/G1 cells.

Hoescht/Fluorescein Diacetate (FDA)/Propidium Iodide (PI)

Method A

1. Label cells with 10 µM Hoechst 33342 for 30 minutes @ 37ºC. (Add 10 µl of 1000 µM stock per ml of media).

2. Harvest cells and resuspend in 400 µl of serum-free media containing 10 µM Hoechst 33342.

3. Add PI to final concentration of 5 µg/ml (Add 2 µl 1 mg/ml stock).

4. Add FDA to final concentration of 0.5 µg/ml (Add 1 µl of 220 µg/ml stock).

5. Incubate at room temperature for 5-15 minutes.

6. Run in flow cytometer.

Method B

1. Harvest cells and resuspend in 400 µl of serum-free media containing 10 µM Hoechst 33342.

2. Label cells with 10 µM Hoechst 33342 for 30 minutes @ 37ºC. (Add 10 µl of 1000 µM stock per ml of media.)

3. Bring to RT. Add PI to final concentration of 5 µg/ml (Add 2 µl 1 mg/ml stock) and FDA to final concentration of 0.5 µg/ml (Add 1 µl of 220 µg/ml stock).

4. Incubate at room temperature for 10-15 minutes.

5. Put on ice and run in flow cytometer.

Stohr M., and Vogt-Schaden M. A new dual staining technique for simultaneous flow cytometric DNA analysis of living and dead cells. In Laerum OD, Lindmo T., Thorud E., (eds), “Flow Cytometry” Vol IV Bergen; Norway: Universitetsforlaget, pp. 96-99 (1979).

Hamori E., Arndt-Jovin D.J., Grimwade B.G., and Jovin T.M., Selection of viable cells with known DNA content. Cytometry 1:132-135. (1980).

Coast Seafood Sample Protocol

Coast Seafood Company sends oyster larvae samples every week to determine whether they are triploid populations. The triploid populations will not reproduce if released into the wild. The following is the protocol used to prepare and analyze the samples. They will always send a diploid sample for you to test.

You will need:

1. 10mg/ml DAPI w/NP40 in 10%DMSO (see Solutions Section)

2. Beaker with ddH₂0 for rinsing spatula

3. Spatula

4. Clean slides

5. 2 tubes per sample

6. funnels

7. 90 mm mesh squares (cut from stock supply;)

1. Label 2 tubes per sample – put a small amount of 1x DAPI into one of the tubes per sample.

2. Using a clean spatula, remove a small amount of larvae from the sample tube.

3. Using a dropper filled with 1x DAPI, drop the sample from the spatula onto a clean slide.

4. Using the flat side of the spatula, smush the larvae until the liquid is a muddy brown color.

5. Using the same dropper, suck up all the liquid and smushed larvae from the slide and place it in the tube containing a small amount of DAPI. You may need a few extra drops of clean DAPI on the slide to get everything. Mix it really well in the tube.

6. Place a funnel and 90 mm mesh square onto the empty tube for the same sample.

7. After mixing really well in the DAPI, suck everything up and push it through the mesh into the empty tube.

8. Before running in the cytometer, syringe and filter each sample. The samples tend to be really concentrated depending on how many larvae you took out at the beginning. You may need to filter twice or add more DAPI if it’s too concentrated.

Reagents used in this protocol and their sources:

DAPI from Accurate Chem. Co.; Cat# 18860 (No Substitutes)

90mm mesh from Small Parts, Inc.; Cat# CMN -0090

Section 10 Protocols for Arrays and RNA preps for Arrays

Basic RNA extraction detailed protocol

Materials needed: Trizol (Life technologies cat #15596-026)

Chloroform

70% ethanol (made with DEPC-treated water)

Qiagen RNeasy spin columns (cat# 74104 mini, cat# 75142 midi)

Buffer RW1 (Qiagen RNeasy kit)

Buffer RPE (with ethanol) (Qiagen RNeasy kit)

RNASE free water (Qiagen RNeasy kit)

Microcentrifuge

Protocol (for 5x10⁶ cells – adjust accordingly):

q Thaw on ice and spin 1000 x g for 5 minutes (in microcentrifuge tube)

q Add 1ml Trizol to pellet (mix by pipetting or with polytron if necessary)
At this step you should freeze on dry ice if you have more than TWO samples.
You can store if you need to at –70°C then thaw at 37°C and proceed with the chloroform step SKIPPING the 5 minutes at room temp.

q Incubate 5’ at room temp

q Add 0.2 ml chloroform

q Shake vigorously

q Incubate at RT for 2-3 minutes

q Centrifuge at 12,000 x g for 2-3 minutes

q Remove aqueous phase (top layer) into a new 1.5 ml microfuge tube

q Slowly add an equal volume of 70% ethanol (RNAse free)

q Load on 2 RNEASY mini columns (no more than 3x10⁶ cells per column)

q Centrifuge at >8,000 x g for 15 seconds

q Discard flow-through

q Add 700 ml buffer RW1 to each column

q Centrifuge at >8,000 x g for 15 seconds

q Discard flow-through and collection tube

q Transfer column to a new collection tube

q Add 500 ml buffer RPE (with ethanol added) to each column

q Centrifuge at >8,000 x g for 15 seconds

q Discard flow-through

q Add 500 ml buffer RPE (with ethanol added) to each column

q Spin at >8,000 x g for 2 min

q Discard flow-through and collection tube

q Transfer each column to a new collection tube

q Centrifuge at full speed for 1 min

q Transfer to a 1.5ml collection tube

q Add 50 ml RNAse-free water to each column

q Centrifuge at > 8,000 x g for 1 minute

q (optional) Add 0.5ml RNASE inhibitor (like RNASE-out)

q Store immediately at –80°C

q Run 2ul on spec and 1ul on Agilent bioanalyzer to access quality. A260/280 should be 1.8-2 and 28s:18s total RNA peak ratio should be 1.4-2.

RNA Extraction from Pancreatic Tissue (LiCl/RNeasy combination method)

Solutions

LiCl/Urea (100ml), pH 5.2

· 12.7g LiCl (3M)

· 36g Urea (6M)

82mg NaOAc (10mM)

20%SDS (100ml)

· 20g SDS

Heat to 50°C to dissolve

PK buffer (prepare fresh at RT) (5ml)

· 20 ml 0.5M EDTA (2 mM final)

· 250 ml 4M NaCl (200mM final)

· 125 ml 20% SDS (0.5% SDS final)

· 50 ml 1M Tris-HCL pH 8.0 (10mM final)

· 100 ml Proteinase K (200 mg/ml final)

· 4455 ml Nuclease-free water

Phenol/Chloroform/IAA (pH <6) – we use Ambion

Chloroform

70% ethanol mixed in DEPC treated water

Qiagen RNeasy mini prep kit

Procedure

q Weigh out 25 mg of RNAlater-stabilized tissue (for approximately 20 ug RNA).

q Homogenize tissue quickly in 1 ml LiCl/Urea buffer on ice with 25ml 20%SDS added. Polytron-type homogenizer (Fisher PowerGen) works well.

q Incubate o/n on ice in cold room (the lithium will precipitate the RNA)

q Centrifuge at 3400rpm (max speed) for 15 min 4°C

q Remove supernatant

q Resuspend in 1 ml LiCl/Urea buffer (without SDS)

q Incubate 30’ on ice

q Centrifuge at 3400 rpm (max speed) for 15 min 4°C

q Remove supernatant

q Resuspend in 500 ml PK buffer and transfer to 1.5 ml Eppendorf tube

q Incubate 20’ @ 37°C

q Add 500 ml phenol/Choloroform/IAA (pH <6)

q Vortex 1 min

q Incubate on ice 5 mins

q Centrifuge at 10,000rpm for 5 min

q Remove upper (aqueous) phase and transfer to new tube

q Add 500 ml phenol/ Chloroform/ IAA (pH <6)

q Repeat above steps

q Remove upper (aqueous) phase and transfer to new tube

q Add 500 ml Chloroform

q Repeat above steps

q Remove upper (aqueous) phase and transfer to new tube

q Add equal volume 70% Ethanol slowly

q Mix gently

q Add immediately to Qiagen RNEASY column (25mg tissue per mini column)

q Centrifuge 15 sec at > 8000 x g

q Discard flow-through

q Add 700 ml Buffer RW1

q Centrifuge 15 sec at > 8000 x g

q Discard flow-through and collection tube & place in new collection tube

q Add 500 ml buffer RPE (with ethanol added)

q Centrifuge 15 sec at > 8000 x g

q Discard flow-through

q Add 500 ml buffer RPE (with ethanol added)

q Centrifuge 2 min at > 8000 x g

q Discard flow-through and collection tube & place in new collection tube

q Centrifuge 2 min at full speed

q Discard collection tube and transfer to a 1.5 ml RNASE-Free tube

q Add 50 ml RNASE-free water directly to the membrane

q Centrifuge 1 min at > 8000 x g

q Add 0.5 ml RNASE-out RNASE inhibitor and store aliquotted at -80°C

q Avoid freeze-thaw cycles

General Instructions for Bioanalyzer RNA

Day before run – Call ahead to make sure the analyzer is available at the time you would like to run. (phone: 5-7285)

Things to take with you

Bucket of ice for samples

Electrode Cleaning Chips (two)

Gel dye matrix mix

Sample Buffer

RNA ladder

Samples

Call ahead (5-7285) to make sure someone can let you in the room.

When you get to the room at Roosevelt

Turn on the heat block (high 95°C) and put RNA ladder in to heat.
Turn on the computer screen.
Turn on Bioanalyzer.
Gel Dye matrix in small box at RT and close to protect from light

Clean Bioanalyzer:

Add 350 ml of RNase ZAP (Ambion Cat. # 9780.9782.9782) to RNase electrode cleaning chip.
Close lid for 1 minute. Remove chip
Add 350 ml of nuclease free water to the water chip.
Close lid for 10 seconds. Remove chip
Air dry 10 seconds.
Set aside chips for use later

Loading Chip.

Put the RNA chip in the “Chip Primer w/syringe”.
Make sure the Chip Primer is set to position C.
Load 9 ml of gel dye matrix mix into the chip in the G well.
Close the top of the chip holder and click shut.
Depress the plunger until it is locked in place and WAIT 30 seconds.
Unlock the plunger, pull the plunger to 1 ml and then release the top.
Add 9 ml to the two other wells labeled G.
The remaining wells are for your samples. You want to load between 100 and 200 ng/well. You may use up to 3 ml of RNA but the remaining volume should be sample buffer provided in the kit. The total volume should be 6 ml. Load the appropriate amount of sample buffer to each well. Then add the appropriate amount of RNA. Make sure you have a plan for adding the RNA so you know which wells are which.
Do NOT leave any sample wells empty. Add 6 ul of sample buffer to any empty wells.
Add 1 ml of the RNA 6000 ladder to the ladder well. This has been heated in the heat block. Then add 5 ml of the Sample Buffer to that same well.
Place the chip on the vortex. Tape it down on the corner. Mix for 1 minute.
Start run within five minutes of loading.

Running chip.

On the computer click on the Agilent 2100 to open the software.
Click on Assay.
Click on Other.
Click on Eukaryotic total RNA or mRNA.
Choose the number of lanes to be analyzed and label them.
Click on OK.
While the samples are running, CLEAN UP the work area so it is clean for the next user.

After run is complete

SAVE the file in the Rabinovitch folder.
Print the electropherograms for your samples.
Clean the Bioanalyzer as described above. When cleaning is complete, flick the liquid from the chip and save the chip for use next time you run an RNA chip
Turn off the Bioanalyzer, heat block, and computer screen.
Gather your things and make sure the door is locked when you leave.
Return the key if necessary.

Make Gel dye mix

Add 400 ml gel matrix + 4 ml dye to a 1.5 ml microfuge tube.
Vortex to mix
Add to provided spin column
Centrifuge @ 7000 x g for 10 min.
Store at 4°C and protect from light

Preparation of cDNA from total RNA (Reverse Transcription, RT)

Description:

This protocol describes the preparation of complementary DNA, cDNA, from total RNA using oligo-dT primers complementary to the poly-A tail of messenger RNAs.  This procedure strongly favors 3’ sequences, a consideration for downstream applications using cDNA prepared in this manner.  However, any reverse primer can be substituted for the oligo-dT primer if desired.  High quality RNA should be used, and it is not necessary to DNAse-treat the RNA prior to cDNA preparation as the initial step in this protocol encompasses a DNAse digestion to remove contaminating DNA.

Step 1

Per rxn: Total RNA (0.1 – 2 mg) x (to 4 ml)

RNAsin (40 U/ml) 0.25 ml (Promega, 2500 U, #N2111)

DTT (100 mM) 0.5 ml

DNAse I (1 U/ml) 0.25 ml (Roche, #776 785)

H₂O (DEPC or RNAse-free) q.s. to 5 ml

Run on RTPCR1, Rabinovitch lab Eppendorf cycler

30’ @ 37°C (This step is to digest residual DNA.)

5’ @ 70°C (This step destroys DNAse and denatures RNA.)

Step 2

Add 1 ml of oligo-dT primer* (0.13 mg/ml) to each tube. Vortex briefly @ 5, spin down.

Run on RTPCR2, Rabinovitch lab Eppendorf cycler

5’ @ 70°C (This step is to denature RNA so that it anneals to primer.)

Snap on ice immediately so RNA doesn’t form secondary structure.

Step 3

To each rxn add: 5x First Strand Buffer 2 ml (Prepare master mix, keep at RT.)

DTT (100 mM) 0.5 ml

dNTPs (10 mM) 0.5 ml

Superscript 0.5 ml (Invitrogen, Biochemistry stores)

Run on RTPCR3, Rabinovitch lab Eppendorf cycler

1 hr @ 45°C (This step is reverse transcription, first strand synthesis.)

When complete, final volume is 9.5 ml. Add 90.5 ml H₂O (q.s. to 100 ml).
Store at –20°C.

*Oligo-dT primer from Roche, #814 270, resuspended in H₂O to 0.13 mg/ml

ENZO RNA prep for generation of cRNA

DAY 1 (FIRST AND SECOND STRAND cDNA SYNTHESIS)

Refer to Affymetrix protocol in blue notebook if you have any questions.

Use program AFFYDNA on eppendorf cycler. Mix and spin down briefly after adding each set of reagents. Use 0.2 ml PCR tube. Most reagents are in “Affy Reagents” box in chest freezer.

Reagent Name	Vol (ul)	Temp	Time	Location
Total RNA (5mg)	10			-80°C
42 uM T7 Oligo dT primer	2.5			-20°C aliquots
		70°C	10 min
5x First Strand cDNA buffer *	4	4°C	HOLD	-20°C stratacooler
0.1M DTT *	2			-20°C stratacooler
10mM DNTP *	1			-20°C aliquots
		42°C	2 min
Superscript II RT	1	4°C	HOLD	-20°C stratacooler
		42°C	1 hour
DEPC H20 *	91	4°C	HOLD	RT (MB grade)
5x Second Strand Rxn Buffer *	30			-20°C
10 mM DNTP *	3			-20°C aliquots
10 U/ml E. coli DNA ligase	1			-20°C
10 U/ml E. coli DNA Polymerase I	4			-20°C
2 U/ml E. coli RNASE H	1			-20°C
		16°C	2h
10 U/ml T4 DNA Polymerase	2	16°C	HOLD	-20°C
		16°C	5 min
0.5 M EDTA	10			RT (MB grade)

* can be made up as a cocktail if doing multiple samples

Store at -20°C if stopping here.

q Transfer to 1.5 –2 ml microfuge tube

q Add 600 ml cDNA binding buffer.

q Vortex 3 seconds. Color should be yellow. If not refer to manual.

q Add 500 ml to cDNA spin column from Affymetrix (Qiagen).

q 1 min @ 8,000xg

q Discard flow-through

q Add remaining mixture cDNA spin column & spin as above

q Transfer column to new collection tube

q Add 750 ml cDNA wash buffer (with ETOH added)

q 1 min @ 8,000xg

q Discard flow-through

q 5 min @ maximum speed (with cap open)

q Transfer column to 1.5 ml collection tube.

q Add 14 ml elution buffer

q Incubate 1 min @ RT

q 1 min @ max speed

q Save 1 ml for analysis later. Be sure to mark as cDNA.

q Store at -20°C

DAY 2 (IN VITRO TRANSCRIPTION & HYB COCKTAIL PREP)

Eppendorf protocol AFFYIVT

Reagent Name	Vol (ml)	Temp	Time	Location
CDNA from above (in 0.2 ml tube)	10	RT		-20°C
Mol bio grade H₂0	12			RT
10x HY Rxn buffer (ENZO #1)	4			-20°C
10x Biotin-labeled ribonucleotides (ENZO #2)	4			-20°C
10x DTT (ENZO #3)	4			-20°C
RNASE inhib (ENZO #4)	4			-20°C
20x T7 RNA pol (ENZO #5)	2			-20°C
		37°C	5h – mix every 30 min

Store @ -70°C if stopping here

q Transfer to 1.5-2 ml tube

q Add 60 ml RNAse-free water to mix and vortex 3 sec

q Add 350 ml IVT cRNA binding buffer & vortex 3 sec

q Add 250 ml ethanol and mix by pipetting

q Apply to IVT cRNA cleanup spin column

q 15 seconds @ 8,000xg

q Transfer to a new collection tube

q Add 500 ml IVT cRNA wash buffer (with ethanol added)

q 15 seconds @ 8,000xg

q Discard flow through

q Add 500 ml 80% ethanol to the spin column

q 15 seconds @ 8,000xg

q Open the cap

q 5 minutes at maximum speed

q Transfer to 1.5 ml tube

q Add 11 ml RNASE free water

q 1 min @ max speed

q add 10 ul RNASE free water

q 1 min @ max speed

q Check spectrophotometer at 1:100 dilution 0.5 ml into 99.5 ml of RNase free water

q Calculate as adjusted cRNA
= measured cRNA – (10/12)*(RNA concentration at start)

q Remove an aliquot (0.5 ml) for Bioanalyzer

q Store at –80°C

Fragmentation

o Plug sample name, measured [cRNA], total RNA into the grey columns of the excel template “template for affy fragmentation.xls”
u:\PUBLIC\AffyPublic\affy public\protocols\ template for affy fragmentation.xls

§ [ cRNA] is calculated using the formula:

adjusted [cRNA]=measured [cRNA] – (10/12)*([total RNA] at start)

§ This will calculate the volume needed (RNA, water, and 5x fragmentation buffer) in the fragmentation reaction for 1,2, or 3 hybridizations.

o Fragment enough RNA for the maximum number of hybridizations possible (if you only have 16 ug, you can only do 1 hybridization. If you have 50 ug, you should fragment enough for 3 hybridizations.)

o 94°C for 35 minutes (PCR program: FRAGMEN). Store on ice or freeze at -80°C

o remove an aliquot (0.5 ul) for Bioanalyzer.

o Adjust the volume of water and cRNA below depending on the volume of the fragmentation reaction (copy from “add to hyb” column on the template).

o Take the aliquots of cRNA before and after fragmentation to the Bioanalyzer to assess quality before sending to CEA.

Hybridization cocktail

Reagent Name	Vol (ml)	Location	Instructions
Fragmented cRNA	13.3	-80°C
B2 oligo	5	-20°C
20x euk hyb control	15	-20°C	Heat to 65°C for 5 min first!!!!!
Herring Sperm DNA	3	-20°C	(10 mg/ml stock)
Acetylated BSA	3	-20°C	(50 mg/ml stock)
2x hyb buffer	150	Cold Room	Discard if yellow
RNASE-FREE H20	110.7	RT

Either place on dry ice and send to CEA immediately or store at -70°C

AFFYmetrix RNA prep for generation of cRNA using IVT kit

DAY 1 (FIRST AND SECOND STRAND cDNA SYNTHESIS)

Refer to Affymetrix protocol in blue notebook if you have any questions.

Use program AFFYDNA on eppendorf cycler. Mix and spin down briefly after adding each set of reagents. Use 0.2 ml PCR tube. Most reagents are in “Affy Reagents” box in chest freezer.

Reagent Name	Vol (ul)	Temp	Time	Location
Total RNA (5mg)	10			-80°C
42 uM T7 Oligo dT primer	2.5			-20°C aliquots
		70°C	10 min
5x First Strand cDNA buffer *	4	4°C	HOLD	-20°C stratacooler
0.1M DTT *	2			-20°C stratacooler
10mM DNTP *	1			-20°C aliquots
		42°C	2 min
Superscript II RT	1	4°C	HOLD	-20°C stratacooler
		42°C	1 hour
DEPC H20 *	91	4°C	HOLD	RT (MB grade)
5x Second Strand Rxn Buffer *	30			-20°C
10 mM DNTP *	3			-20°C aliquots
10 U/ml E. coli DNA ligase	1			-20°C
10 U/ml E. coli DNA Polymerase I	4			-20°C
2 U/ml E. coli RNASE H	1			-20°C
		16°C	2h
10 U/ml T4 DNA Polymerase	2	16°C	HOLD	-20°C
		16°C	5 min
0.5 M EDTA	10			RT (MB grade)

* can be made up as a cocktail if doing multiple samples

Store at -20°C if stopping here.

q Transfer to 1.5 –2 ml microfuge tube

q Add 600 ml cDNA binding buffer.

q Vortex 3 seconds. Color should be yellow. If not refer to manual.

q Add 500 ml to cDNA spin column from Affymetrix (Qiagen).

q 1 min @ 8,000xg

q Discard flow-through

q Add remaining mixture cDNA spin column & spin as above

q Transfer column to new collection tube

q Add 750 ml cDNA wash buffer (with ETOH added)

q 1 min @ 8,000xg

q Discard flow-through

q 5 min @ maximum speed (with cap open)

q Transfer column to 1.5 ml collection tube.

q Add 14 ml elution buffer

q Incubate 1 min @ RT

q 1 min @ max speed

q Save 1 ml for analysis later. Be sure to mark as cDNA.

q Store at -20°C

Overnight (IN VITRO TRANSCRIPTION & HYB COCKTAIL PREP)

Thaw all components and do NOT assemble on ice because 10X IVT Labeling buffer has spermidine that will precipitate the cDNA

Reagent Name	Vol (ml)	Temp	Time	Location
CDNA from above (in 0.2 ml tube)	6	RT		-20°C
Mol bio grade H₂0	14			RT
10x IVT Labeling buffer	4			-20°C
IVT Labeling NTP Mix	12			-20°C
IVT Labeling Enzyme mix	4			-20°C
		37°C	16 hours

Store @ -70°C if stopping here

q Transfer to 1.5-2 ml tube

q Add 60 ml RNAse-free water to mix and vortex 3 sec

q Add 350 ml IVT cRNA binding buffer & vortex 3 sec

q Add 250 ml ethanol and mix by pipetting

q Apply to IVT cRNA cleanup spin column

q 15 seconds @ 8,000xg

q Transfer to a new collection tube

q Add 500 ml IVT cRNA wash buffer (with ethanol added)

q 15 seconds @ 8,000xg

q Discard flow through

q Add 500 ml 80% ethanol to the spin column

q 15 seconds @ 8,000xg

q Open the cap

q 5 minutes at maximum speed

q Transfer to 1.5 ml tube

q Add 11 ml RNASE free water

q 1 min @ max speed

q add 10 ul RNASE free water

q 1 min @ max speed

q Check spectrophotometer at 1:100 dilution 0.5 ml into 99.5 ml of RNase free water

q Calculate as adjusted cRNA
= measured cRNA – (10/12)*(RNA concentration at start)

q Remove an aliquot (0.5 ml) for Bioanalyzer

q Store at –80°C

Fragmentation

§ [ cRNA] is calculated using the formula:

adjusted [cRNA]=measured [cRNA] – (10/12)*([total RNA] at start)

§ This will calculate the volume needed (RNA, water, and 5x fragmentation buffer) in the fragmentation reaction for 1,2, or 3 hybridizations.

o Fragment enough RNA for the maximum number of hybridizations possible (if you only have 16 ug, you can only do 1 hybridization. If you have 50 ug, you should fragment enough for 3 hybridizations.)

o 94°C for 35 minutes (PCR program: FRAGMEN). Store on ice or freeze at -80°C

o remove an aliquot (0.5 ul) for Bioanalyzer.

o Adjust the volume of water and cRNA below depending on the volume of the fragmentation reaction (copy from “add to hyb” column on the template).

o Take the aliquots of cRNA before and after fragmentation to the Bioanalyzer to assess quality before sending to CEA.

Hybridization cocktail

Reagent Name	Vol (ml)	Location	Instructions
Fragmented cRNA	15	-80°C
B2 oligo	5	-20°C
20x euk hyb control	15	-20°C	Heat to 65°C for 5 min first!!!!!
Herring Sperm DNA	3	-20°C	(10 mg/ml stock)
Acetylated BSA	3	-20°C	(50 mg/ml stock)
2x hyb buffer	150	Cold Room	Discard if yellow
DMSO	30
RNASE-FREE H20	79	RT

Either place on dry ice and send to CEA immediately or store at -70°C
MCAT Genotyping Protocols

(from John Morton in Warren Ladiges lab)

Description:

The 5’ Primer OCT/CAT, located in the OCT leader sequence, can be used with three different 3’ primers giving three primer sets which amplify products of different size from the MCAT transgene. OCT/CAT and 3’ primer CAT1 amplify a 692 bp product, OCT/CAT and 3’ primer CAT2 amplify a 900 bp product, and OCT/CAT and 3’ primer CAT3 amplify a 1429 bp product.

The primer CATX7 and CATX8 span exon 7 and 8 of the human catalase gene. These primers can be used to genotype cDNA prepared from total RNA using oligo dT primers, a key advantage over using the OCT/CAT Forward primers with on the the three Reverse primers used for genotypeing DNA This is because the OCT/CAT Forward primers bind the 5’ end of the gene, and oligo dT primed RT-PCT strongly favors 3’ sequences. Expected product size using the CAT X7 and CATX8 primers is 256 bp.

Primers (purple rack in fridge #2):

OCT/CAT 5’- ctg agg atc ctg tta aac aat gc – 3’

CAT2 5’– ggg aaa gtc tcg ccg cat ctt c – 3’

Primers (located in –20 C freezer FNH Box 10A)

CATX7 (hCAT F) 5’- gaa gat gcg gcg aga ctt tcc c –3’

CATX8 (hCAT R) 5’- cta tct gtt caa cct cag caa ag –3’

These primers are not in the lab but can be obtained from John Morton.

CAT1 5’ – gaa gtc cca gac cat gtc cgg at – 3’

CAT3 5’ – ctc cgc act tct cca gaa tat tg – 3’

Reaction Mix for OCT/CAT & CAT2 Reaction Mix for CATX7 & CATX8
(For one 25ul reaction): (For one 25ul reaction):

H₂O 12.5ml 10X PCR Buffer +Mg²⁺ 2.5ml

10X PCR Buffer 2.5ml 20mM 5’Primer 1.0ml

25mM MgCl2 1.5ml 20mM 3’Primer 1.0ml

25mM 5’Primer 1.0ml 10mM dNTP 0.5ml

25mM 3’Primer 1.0ml Taq Polymerase 0.2ml

25mM dNTP 0.2ml cDNA template 5.0ml

5X Rapid-Load Buffer* 5.0ml H₂O 14.8ml
(optional -- add 5 ml H₂0 if not available)

Taq Polymerase 0.3ml

DNA 1.0ml

*= 5X R.L. Buffer – OriGene Technologies, Catalog # RL-105

Thermocycler Settings for Thermocycler Settings for
OCT/CAT & CAT2: CATX7 & CATX8

94°C for 4 min 94°C for 4 min

cycle 30 times cycle 30 times

94°C for 30 sec 94°C for 30 sec

55°C for 30 sec 60°C for 30 sec

72°C for 1.5 min 72°C for 1.5 min

72°C for 5 min 72°C for 5 min

4°C forever 4°C forever

Run products on a 1% agarose gel. Run products on a 2% agarose gel

Section 11 Macros, SAM, GeneTraffic hints

Entropy Macro

Written by Mike Shen April 2004

Theory: ENTROPY (Diversity)

A measure or variation or diversity defined on the probability distribution of observed events. Specifically, if P is the probability of an event a, the entropy H(A) for all events a in A is:

H(A) = -SUM_a P(a) log₂ P(a)

i.e., The sum of the log2 probs of the model p, weighted by the real probs p.

The quantity is zero when all events are of the same kind, p =1 for any one a of A and is positive otherwise. Its upper limit is log_2 N where N is the number of categories available (see degrees of freedom) and the distribution is uniform over these, p = 1/N for all a of A (see variety, uncertainty, negentropy). The statistical entropy measure is the most basic measure of information theory.

Simpson's index of diversity

Simpson's index is the first of a set of non-parametric approaches to determining sample heterogeneity. Simpson (1949) did not want to make assumptions about the distribution of the species abundance curve and so defined the following:

Simpson's index

Simpson index formula

where D is Simpson's index and can be defined as the probability of two individuals in a random sample being in same category. Pi is the proportion of category i in the community.

Simpson's index of diversity = 1 - D ,

Range(0 to ~1),

Maximum (1 - 1/categories).

The probability that two randomly selected individuals in a community are of the different categories.

Also see http://www.tiem.utk.edu/~gross/bioed/bealsmodules/simpsonDI.html

Entropy Macro “SID.xlm” (general purpose)

(1) Excel Macro is available on U:\public\Entropy_Diaversity Macro\SID.xlm

(2) Example data is in U:\public\Entropy_Diaversity Macro\tsSID.xls with following format:

a. Title: starts at row #5

b. Data Set: First set starts at row #6 with next data set at next row.

c. Data entry: First event starts at column C with next event at next column.

Example (tsSID.xls):

	A	B	C	D	E	FISH abnormality template	G	H	I	J	K	L	M	N	O	P	Q	R	S	T	U	V	W	X	Y	Z	AA	AB	AC	AD	AE	AF	AG
2						FISH abnormality template
3
4
5			0r1g	0r2g	0r3g	0r4g	1r0g	1r1g	1r2g	1r3g	1r4g	2r0g	2r1g	2r2g	2r3g	2r4g	2r5g	3r0g	3r1g	3r2g	3r3g	3r4g	3r5g	4r0g	4r1g	4r2g	4r3g	4r4g	5r2g	5r3g	5r4g	5r5g	Cell #
6	121 ja20 chrom 17		0	0	0	0	0	1		0	0	0	5	#	6	0	0	0	0	31	9	0	0	0	0	0	1	1	0	0	0	0	199
7	12u #12 chr8		0	0	0	0	0	2		1	0	0	1	85	5	0	0	0	1	3	0	0	0	0	0	0	0	1	0	0	0	0	103
8	131j d6 (2d=4)		0	0	0	0	0	0		0	0	0	3	#	5	1	0	0	1	#	3	0	0	0	0	0	0	0	0	0	0	0	204
9	27u #17 chr 8		0	0	0	0	0	5		2	0	0	5	95	#	0	0	0	2	11	2	0	0	0	0	0	0	1	0	0	0	0	145
10	35r 5cm		0	0	0	0	0	0		0	0	0	0	#	6	1	0	0	0	1	0	0	0	0	0	0	0	0	0	0	0	0	158
11	36x 80 cm		0	0	0	0	0	1		0	0	0	2	#	3	0	0	0	0	4	1	0	0	0	0	0	0	0	0	0	0	0	198
12	37u 15cm		0	0	0	0	0	2		0	0	0	1	#	6	0	0	0	0	3	4	0	0	0	0	0	0	0	0	0	0	0	210
13	46t #21 chr 18		0	0	0	0	0	1		1	0	0	1	#	5	0	0	0	1	4	1	0	0	0	0	0	0	1	0	0	0	0	171
14	46t 21 cm		0	0	0	0	0	3		1	0	0	2	#	8	1	0	0	0	2	2	0	0	0	0	0	0	0	0	0	0	0	200
15	4z c23 ch17		0	0	0	0	0	5		3	0	0	5	#	#	2	0	0	2	12	3	1	0	0	0	1	0	2	0	0	1	0	200
16	82u #2 chr 8		0	0	0	0	0	2		0	0	0	2	#	3	0	0	0	0	5	0	0	0	0	0	0	0	2	0	0	0	0	201
17	95u #22 chr 8		0	0	0	0	0	2		1	0	0	2	85	8	2	0	0	1	0	3	0	0	0	0	0	0	0	0	0	0	0	104

Column A and B are reserved for Label.

Column C is the first event, column D is the second event, etc up to column AF. The entry is the frequency or the probability.

Row 1-4 is for general labels.

Row 5 is the detail labels

Row 6 is the first data set, row 7 is the second data set, etc.

To Run SID.xlm

(1) prepare the excel data as shown in example above.

(2) Click on SID.xlm to open the macro.

(3) Click on Tools\Macro\Macros (or ALT F8) to bring up function list of this macro. Click Start_SID and RUN

(4) At prompt, choose your excel data file.

(5) Macro takes over and summary is appended at completion:

Result summary (Output by macro):

	AP	AQ	AR	AS	AT	AU
2
3
4
5	TotalSid	TotalEntropy	ArmSid	ArmEntropy	CentomereSid	CentomereEntropy
6	0.46	1.46	0.37	0.94	0.21	0.64
7	0.31	1.14	0.21	0.67	0.20	0.63
8	0.34	1.05	0.26	0.65	0.12	0.42
9	0.55	1.88	0.36	1.00	0.36	0.99
10	0.14	0.51	0.06	0.23	0.09	0.29
11	0.18	0.67	0.13	0.44	0.07	0.26
12	0.16	0.63	0.10	0.35	0.12	0.38
13	0.21	0.81	0.14	0.49	0.12	0.42
14	0.19	0.73	0.09	0.31	0.16	0.52
15	0.52	1.89	0.36	1.06	0.32	0.96
16	0.16	0.63	0.11	0.41	0.09	0.33
17	0.32	1.13	0.13	0.42	0.32	0.92

Entropy Macro “SID.xlm” (Telomere arm gain_loss, centromere gain_loss)

(1) Excel Macro is available on U:\public\Entropy_Diaversity Macro\SID.xlm

(2) Example data is in U:\public\Entropy_Diaversity Macro\telomere_SID.xls: There are more calculations that are associated with telomere data that is not applicable in general. Use this file as your template. Rename the file and edit the content, save the file before running the macro.

To Run SID.xlm

(1) prepare the excel data as shown in example above.

(2) Click on SID.xlm to open the macro.

(3) Click on Tools\Macro\Macros (or ALT F8) to bring up function list of this macro. Click Start_SID and RUN

(4) At prompt, choose your excel data file.

(5) Macro takes over and summary is appended at completion:

Result summary (Output by macro):

	AP	AQ	AR	AS	AT	AU
2
3
4
5	TotalSid	TotalEntropy	ArmSid	ArmEntropy	CentomereSid	CentomereEntropy
6	0.46	1.46	0.37	0.94	0.21	0.64
7	0.31	1.14	0.21	0.67	0.20	0.63
8	0.34	1.05	0.26	0.65	0.12	0.42
9	0.55	1.88	0.36	1.00	0.36	0.99
10	0.14	0.51	0.06	0.23	0.09	0.29
11	0.18	0.67	0.13	0.44	0.07	0.26
12	0.16	0.63	0.10	0.35	0.12	0.38
13	0.21	0.81	0.14	0.49	0.12	0.42
14	0.19	0.73	0.09	0.31	0.16	0.52
15	0.52	1.89	0.36	1.06	0.32	0.96
16	0.16	0.63	0.11	0.41	0.09	0.33
17	0.32	1.13	0.13	0.42	0.32	0.92

Result associated with telomere:

	AH	AI	AJ	AK	AL	AM	AN
2
3
4
5	% normal	% arm loss	% arm gain	% arm abnormal	%. Cen. Loss	%. Cen. Gain	% Cen. Abnl
6	71.36	2.01	21.11	23.12	3.02	8.54	11.56
7	82.52	6.80	4.85	11.65	3.88	6.80	10.68
8	80.39	0.49	14.71	15.20	1.96	4.41	6.37
9	65.52	10.34	11.03	21.38	8.28	13.10	21.38
10	92.41	2.53	0.63	3.16	0.00	4.43	4.43
11	90.40	4.55	2.53	7.07	1.52	2.02	3.54
12	91.43	1.90	3.33	5.24	1.43	4.76	6.19
13	88.89	3.51	4.09	7.60	1.75	4.68	6.43
14	90.00	2.50	2.00	4.50	2.50	6.00	8.50
15	68.50	10.50	11.00	21.50	6.00	12.50	18.50
16	91.54	2.49	3.48	5.97	1.99	2.49	4.48
17	81.73	2.88	3.85	6.73	4.81	13.46	18.27

Where

% normal = #2r2g/total

% arm gain = #xrng/total where x>2

% arm loss = #xrng/total where x<2

% cen gain = #nryg/total where y>2

% cem loss = #nryg/total where y<2

Transfer of Data from CEA to GeneTraffic

Transfer of array data from the CEA to the U drive

Down loading of array data into Gene Traffic

Transfer of array data from the CEA to the U drive

When the arrays have been scanned the data is stored on the RA at the CEA.
To access this data you will need to open “SSH secure ftp” usually on your desktop. Click on quick connect and fill in the boxes

ra.microslu.washington.edu

petersr

Password

Type our password in the box (you can get this from Katy) and click OK.

Operations

The page will then look like this.

On the left side go to

My Computer

Lab_Rabinovitch on ‘Pathology\Files’ (U:)

PUBLIC

Affy Public

AFFY RAW DATA FROM CEA

Create a folder for your data

Now go to the right side

Click on the folder and then files you want to transfer, include *.CEL, *.CHP and *.DAT

Click on “Operations” in the toolbar and then download.

Down loading of array data into Gene Traffic

· Open Iobion Informatics GeneTraffic

· http://128.208.135.63/?

· Log on with password provided by Dick Byer dbeyer@u.washington.edu

· Click on “Create New Project”

SAM Autopilot macro

Purpose:

The SAM Autopilot macro is intended to automate some of the common ways of producing gene lists and probe sets using SAM in excel. Starting with a gene table or probe set, it generates .txt files containing gene lists and probe sets for roughly 85, 180, and 280 of the most statistically significant genes.

Installation:

SAM Autopilot is an excel add-in. To install it, select the tools–>Add-Ins menu item in excel. When the Add-Ins dialog box appears, push the Browse button and navigate to U:\PUBLIC\Affy Public\Excel Macros. There should be one or more files named SAM Autopilot <date>; chose the one with the most recent date. When it asks if you would like to copy it to your AddIns director, click yes.

Creating a Toolbar button for SAM Autopilot (optional):

If you’d like to add a menu item or toolbar button for SAM Autopilot, you can do the following: First, choose the Tools->Customize menu item. In the dialog that appears, chose the Commands tab and select the Macro item from the categories list. In the commands list you should see two items, “Custom Menu Item” and “Custom Button.” Drag and drop the “custom button” into the toolbar just to the right of the “SAM Plot Control” button. Without closing the dialog box, click on the new toolbar button. A black border should highlight it, and the Modify Selection button in the dialog box should now be enabled. Click on the Modify Selection button, and choose Assign Macro. In the Assign Macro dialog box, type “GenerateSAMFiles”, then click OK. Click on the Modify Selection button again, select Name, and type “SAM Autopilot”. Click on Modify Selection one more time, and then select “Text Only (Always)”. Finally, click on the close button to close the dialog box.

Uninstallation:

Select the tools–>Add-Ins menu item in excel. When the Add-Ins dialog box appears, uncheck the checkbox next to SAM Autopilot in the add-ins list.

Using SAM Autopilot:

To use SAM Autopilot, open the file containing the gene table. If this is the first time you’ve run SAM Autopilot on the data, make sure that the worksheet with the gene table is the one in front. Next, run the macro by pressing control-T (or press the toolbar button if you created one).

If this is the first time you’ve run SAM Autopilot, a dialog box will appear stating:

“Data categories not found - please enter the data categories, then run the macro again.” The active sheet will now be the “Categories” sheet. In the first row is a list of the column names from the gene table. In the second row is the SAM category for each column – initially these will all be set to “1”. You will also see a “Type” field (initially “One Class”) and a suffix field (initially blank).

The categories can be assigned according to the standard SAM types (1 or 2 for two class, 1 to n for multiclass, -n to n for paired, etc.). If a category field is left blank, the macro will ignore that chip in the SAM analysis – this is an easy way of analyzing a subset of the data (for example comparing just young wild type vs young KO). The type field can be any one of the SAM analysis types, but generally you won’t need to change it (by default is figures it out the analysis type based on what you entered for categories). The suffix field is a suffix to be added to all of the file names, for example “YWT vs YKO”; it’s primarily intended to be used when you’re analyzing subsets of the data and don’t want to overwrite the files you’ve generated from other subsets.

After adjusting the categories, you can then run the SAM Autopilot macro again (using ctrl-T or the toolbar button). IMPORTANT: Do not switch to another application while SAM Autopilot is running. SAM Autopilot controls SAM by emulating a user at the keyboard; if you switch to another application those key presses will go to the other application instead of SAM. (The macro certainly won’t work, and who knows what the other application might do).

While the macro runs, you will see the SAM plot controller dialog box once or twice (roughly once for every 30,000 genes in the analysis). When you see the SAM plot dialog, just press “List Delta Table”. (If you hit “list significant genes” or adjust the slider, don’t worry – the data will still be good, but the macro will take a few seconds longer to run). Since the current version of SAM can’t handle more than 32,000 genes, larger gene sets are broken up into smaller sets of 30000 genes.

When the macro has finished, it will have created a new folder in the same folder as the original gene table. The folder will be called <gene table filename>[-<suffix>] (for example “GRKO-YWT vs YKO”), and will contain a list of probe sets and gene lists. The files will have the names <gene table filename>-<list type>[-suffix]-<set size (1, 2, or 3)><gene subset>.txt (for example “GRKO-ProbeSet-YWT vs YKO-2A.txt”). The list type is either genelist or probeset, the set size is 1 for ~85 genes, 2 for ~180 genes, and 3 for ~280 genes. The gene subset is only relevant for analyzing more that 30000 genes - files generated from the analysis of the first 30000 genes will end in ‘A’, the next block of genes in ‘B’, etc.. The new gene lists and probe sets can be imported into gene traffic as tab-delimited text files.

In addition to the new files, SAM Autopilot also saves the gene lists, probe sets, and SAM output as new worksheets in the current workbook. The SAM Output[-suffix]-<setsize><gene subset> sheets in particular provides a convenient record of the genes and category assignments used to generate a particular gene list.

Extracting pixel intensity data from BAC array images

using GenePix software

FHCRC computer lab is located in CD-111; use computers #15, 16, or 17

To logon to computer, user: affychip

Password: DNA_Array

1. Copy GAL file onto the desktop—this contains positional information for each clone. Note: There are several files so compare slide # and batch # to be sure you copy the right file.

To access GAL files, open Internet Explorer and go to this website: ftp://milano.fhcrc.org/spotted.arrays/array/files/Human_BAC/GAL_Files/

2. Open GenePixPro3.0 (shortcut on desktop).

3. Check the ratio setting 635nm/532nm (upper left corner).

4. Open image file. (Click on button with picture of file folder—upper right)

To confirm that the original (unmodified) image is shown—equalize brightness & contrast settings.

5. Open the GAL list and move it with mouse to align it roughly with the blocks.

6. Click on the zoom feature (magnifying glass with “+” in the center) and select 4 blocks with marquee.

7. Click on feature and block alignment button (button with circle) and select “find all blocks and align features”.

8. Confirm the feature alignment by scrolling over each block—adjust as necessary using block manipulation feature (square with arrow pointing to upper left corner).

9. Manipulate features using the single feature button (circle with arrow pointing to upper left and away).

a. All absent features should already be marked with null symbols.

b. Bad features or those not clearly defined should be marked as “Absent”—hit A.

c. To shrink the feature size, hit CTRL and down arrow.

d. To enlarge the feature size, hit CTRL and up arrow.

e. Any regions with scratches should be marked to exclude those samples.

Note: Background pixels are calculated outside of a 2-pixel exclusion region surrounding each feature.

10. Click “ANALYZE” button (upper right).

11. Save gpr result file. Use “SAVE AS” selection in upper left corner.

12. Transfer files to memory stick or CD.

Analysis of BAC array data using R scripts

Note: This analysis uses BATCH mode in the command line not in the R program.

1. Open command line for BATCH analysis in R.

Go to START menu, then to Run. Open “cmd”

The command window will open and the “M:>” prompt will appear.

2. Type “cd BAC files” to change directory to BAC files.

3. Open normalization text file and verify/edit lines highlighted below. The rest of the file can be left as is.

## R script to normalize BAC arrays

############## BEGIN USER SPECIFICATION OF INFO NEEDED BY PROGRAM ##############

## Name of file containing map info: ID, Block, Column, Row, Chrom, Mid, Start, End

## for *ALL* BACs (if locations not known then missing values are present)

## (this file is created by another program)

map.file <- "map10K_OG3_05-21-04.txt" # New map

#map.file <- "map10K_11-17-03.txt"

## File containing list of BACs to drop due to FISH results

#bad.FISH.file <- "259CloneSamples_ExcludeOnFISH_111203.txt"

bad.FISH.file <- "Exclude_On_FISH_04-15-04.txt"

## File containing list of BACs to drop due to performance issues

poor.BAC.file <- "BAD_BACs_06-09-03.txt"

## List of clones that have "bad" FISH results but that are still of interest

bad.FISH.to.use <- c("", "")

# Specify name of directory containing GPR files and names of GPR files.

# 'dir' can be full path or relative to directory in which this script is run.

dir <- "GPR files"

files <- c("2004-06-18_BAC-709_XXRef-vs-39Y-PAL.gpr",

"2004-06-18_BAC-710_XXRef-vs-39Y-NAL.gpr",

"2004-06-18_BAC-711_XXBulk-vs-39Y-Bulk.gpr",

"2004-06-10_BAC-706_XXRef-vs-39Y-P_0500.gpr",

"2004-06-10_BAC-708_XXRef-vs-39Y-N_0500.gpr",

"2004-05-14_BAC-599_XXRef-vs-39YCrypt.gpr"

)

#files <- paste(names,".gpr",sep="")

## Specify names of files that raw and normalized spot data should be written to

outfile.raw <- "Samples-20040621-raw.txt" #paste(names,"-raw.txt", sep="") #"Samples_ABCDE_raw-spot.txt"

outfile.norm <- "Samples-20040621-norm.txt" #paste(names,"-norm.txt", sep="") #"Samples_ABCDE_norm-spot.txt"

## Names to be used in labelling samples in 'outfile.raw' and 'outfile.norm'

sample.names <- c("39Y-PAL", "39Y-NAL", "39Y-bulk", "39Y-P", "39Y-N", "39Y-crypt")

## Should normalization also include scale adjustment (i.e. should we scale

## values so each array has the same variability as the others)? TRUE/FALSE

scale.adjust <- FALSE

## *IF* scale.adjust == TRUE then should adjustment be done using the robust

## MAD (median absolute deviation)? TRUE/FALSE (otherwise use variance)

use.mad <- FALSE

## If you wish to set the scale of the arrays to a particular value then set

## 'specify.scale' to TRUE, and set 'scale.val' to the value you want to use.

## Otherwise set 'specify.scale' to FALSE, and scale.val to anything (its

## value will be ignored). If a value is not specified, then the value used

## will be the median value of the arrays being normalized (those in 'files')

specify.scale <- FALSE

scale.val <- NULL

## This code/script/program assumes that the "reference" sample has been

## labelled with Cy3 (green) dye and the "test" sample with Cy5 (Red) dye.

## The log-ratios are computed (by default) as log2('Cy5 sample'/'Cy3 sample').

## However, if we want log2(test/reference) and the reference is not labelled

## with Cy3 then we won't get what we want by default. The variable 'ref.Cy5'

## allows us to change the default and to calculate the log-ratio as

## log2(Cy3/Cy5) instead. 'ref.Cy5' should contain the position(s) in the vector

## 'files' of all experiments for which the reference was labelled with Cy5

## rather than Cy3. For example a value of c(2,4) would indicate that the

## 2nd and 4th files names in 'files' have Cy5 labelled references. If there

## are no such files then ref.Cy5 should have value c() or NULL.

ref.Cy5 <- c()

## Did any of the reference samples contain >=1 "Y" chromosome? TRUE/FALSE

any.Y.refs <- TRUE

## Which files in 'files' used a reference sample having a "Y" chromosome. This

## is only used if 'any.Y.refs == TRUE, and should be a vector of position

## numbers (e.g. c(1,4) means 1st and 4th file have ref. with Y chrom.)

Y.chrom.samps <- 1:length(files)

## Should code (following user specified values) be printed in the BATCH output

## file, or just values/messages generated by the program? TRUE/FALSE

show.code <- FALSE

########\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\#########

######## END USER SPECIFICATION: NO CHANGES BEYOND THIS POINT!!! #########

########/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/#########

options(echo=show.code)

cat("\n\n\n")

## Add directory info only file names to create full path to files

dfiles <- paste(dir,files,sep="/")

## CHECK FOR USER SPECIFICATION ERRORS ###

## read in and run error checking code

source("norm_input_errors.r")

###### BEGIN PREP OF BAC MAP ########

# read in map info (BAD BACs already dropped), and keep subset of columns

map.info <- read.table(map.file, header=T, sep="\t", as.is=T)

map.info <- map.info[,c("ID","Block","Column","Row","Chrom","Mid","Start","End")]

## delete BACs not having location info

map.info <- map.info[!is.na(map.info$Mid),]

## read in BACs from bad FISH file

bad.FISH <- scan(bad.FISH.file, what="")

## read in BACs from poor performance file

poor.BAC <- scan(poor.BAC.file,what="")

## make BAC names uppercase (in case they aren't) and combine

## them into a single vector

BACs.to.drop <- unique(toupper(c(bad.FISH, poor.BAC)))

## Remove some BACs from list to examine

BACs.to.drop <- BACs.to.drop[!(BACs.to.drop %in% bad.FISH.to.use)]

map.info <- map.info[!(map.info$ID %in% BACs.to.drop),]

###### END PREP OF BAC MAP ########

############# START LOOPING OVER ARRAYS ##############

## read data into a single data frame

for (indx in 1:length(dfiles)) {

v <- substring(read.table(dfiles[indx],as.is=T,skip=2,nrows=1),22,24)

if (v == "1.4") skip <- 30 #how many lines to skip in the header

chip <- read.table(dfiles[indx], sep="\t", header=T, skip=skip, as.is=T,

na.strings="Error", comment.char="", check.names=FALSE)

# subset out columns, then rename two fields

chip <- chip[,c("Block","Column","Row","F635 Median","F532 Median","Flags")]

names(chip)[4:5] <- c("R","G")

## ******** add map info to BAC data ***********

## add chromosome and location info, then drop some merging variables

chip <- merge(map.info,chip,by=c("Block","Column","Row"),sort=FALSE)

chip <- chip[,!(names(chip) %in% c("Column","Row"))]

## ******** End adding map info ***********

## Save BAC name and location info in a dataframe, so we can drop them from 'chip'

## Also create dataframes for storing raw and normalized data

if (indx == 1) {

info.df <- chip[,c("ID","Chrom","Mid","Start","End")]

raw.df <- as.data.frame(matrix(NA,nrow=nrow(chip),ncol=2*length(dfiles)))

norm.df <- as.data.frame(matrix(NA,nrow=nrow(chip),ncol=length(dfiles)))

}

chip <- chip[,!(names(chip) %in% c("ID","Chrom","Mid","Start","End"))]

## calculate raw data log-ratios and intensity values for each spot

chip <- transform(chip, log.ratio=log2(R)-log2(G), intensity=.5*(log2(R)+log2(G)))

## Change log-ratios to log2(G/R) for experiments where reference labelled Red

if ( indx %in% ref.Cy5 ) chip[,"log.ratio"] <- (-1)*chip[,"log.ratio"]

######## **** ASSIGN MISSING VALUES TO LOG RATIOS ***** ############

## insert missing values for log.ratios of flagged spots

chip$log.ratio[chip$Flags<0] <- NA

## insert NAs for log.ratios of Y chrom BACs when ref. sample has no Y chrom

if (!any.Y.refs || !(indx %in% Y.chrom.samps))

chip$log.ratio[which(info.df$Chrom=="Y")] <- NA

## drop more un-needed variables

chip <- chip[,!(names(chip) %in% c("R","G","Flags"))]

######## **** END SETTING LOG RATIOS TO MISSING ***** ############

############### ***** NORMALIZE DATA ****** #################

## create vector to store normalized ratios in

norm.ratio <- rep(NA, nrow(chip))

## make vector indicationg rows that don't have log ratios of NA

not.excluded <- !is.na(chip$log.ratio)

for(k in 1:length(unique(chip$Block))) {

blk <- (chip$Block == k & not.excluded)

## Get loess fit

pt.fit <- predict(loess(log.ratio~intensity, data=chip, subset=blk,

degree=1, span=.65, family="symmetric",

control=loess.control(surface="direct")),

newdata=data.frame(chip$intensity[blk]))

## get normalized value by subtracting fitted value from raw value

norm.ratio[blk] <- chip$log.ratio[blk] - pt.fit

}

raw.df[, 2*(indx-1) + 1:2] <- chip[,c("log.ratio","intensity")]

norm.df[,indx] <- norm.ratio

}

## Do scale adjustment if chosen by user

if (scale.adjust) {

if (use.mad) {

var.fun <- mad

var.meth <- "Median Absolute Deviation (MAD)"

abbrev<- "MAD"

} else {

var.fun <- sd

var.meth <- "Standard Deviation (SD)"

abbrev<- "SD"

}

array.scale.vals <- round(apply(norm.df, 2, var.fun, na.rm=TRUE),dig=3)

if (!specify.scale) scale.val <- round(median(array.scale.vals),dig=3)

norm.df[] <- apply(norm.df, 2, function(x,sv,vf) {

x * (sv / vf(x,na.rm=T)) }, scale.val, var.fun)

cat("\n\n*****************************************************************************\n")

cat("* Applied scale normalization to all arrays.",

"\n* The measure of variability used was the: ", var.meth,

"\n* Arrays were scaled to have a",abbrev," of:", scale.val,"\n",

"\n* The individual arrays had the following original scale values:",

"\n* -------------------------------------------------------------\n")

print(data.frame(Sample=I(sample.names), Scale.Value=array.scale.vals))

cat("*****************************************************************************\n\n\n")

}

## create data.frame to output to file, then reorder columns

names(norm.df) <- sample.names

raw.names <- paste(rep(sample.names,each=2),c("lr","int"),sep=".")

names(raw.df) <- raw.names

norm.df <- cbind(info.df, norm.df)

raw.df <- cbind(info.df, raw.df)

## sort rows into order by Chrom, Mid and ID

norm.df <- norm.df[order(norm.df$Chrom, norm.df$Mid, norm.df$ID),]

raw.df <- raw.df[order(raw.df$Chrom, raw.df$Mid, raw.df$ID),]

## Output raw and normalized data to files

write.table(raw.df, outfile.raw, sep="\t", row.names=F)

write.table(norm.df, outfile.norm, sep="\t", row.names=F)

options(echo=TRUE)

print(proc.time())

q(save="no")

4. Run script to normalize data. To run analysis, type

M:\BAC files> “c:\ProgramFiles\R\rw1090\bin\rcmd” BATCH normalization-lisa-3.txt test.txt

5. Run script to average pixel intensities (each BAC is spotted twice per array). Adjust highlighted portions as required.

## This file reads in normalized data files and creates a file

## with only one row per BAC, i.e. it averages over all the spots

## for a BAC to produce one value for each BAC.

## File containing normalized spot data

spot.file <- "Samples-20040621-norm.txt"

## Name of file to contain normalized data averaged by BAC (i.e. one value per BAC)

outfile.ave <- "Samples-20040621-norm-ave.txt"

## drop BACs with only one good spot (of the two on the array)

drop.single.spots <- TRUE

spots <- read.table(spot.file, sep="\t", header=T, as.is=T)

uniq.chip <- unique(spots[,c("ID","Chrom","Mid","Start","End")])

n.samps <- ncol(spots) - ncol(uniq.chip)

if (n.samps >1) {

aves <- t(apply(as.matrix(uniq.chip$ID),1,function(x,id,tmp) {

ind <- (id == x)

cm <- colMeans(tmp[ind,],na.rm=TRUE)

if (drop.single.spots) cm[colSums(is.na(tmp[ind,])) >= 1 ] <- NA

},spots$ID, spots[,!(names(spots) %in% names(uniq.chip))]))

} else {

aves <- apply(as.matrix(uniq.chip$ID),1,function(x,id,ns,tmp) {

ind <- (id == x)

cm <- mean(tmp[ind],na.rm=TRUE)

if (drop.single.spots) cm[sum(is.na(tmp[ind])) >= 1 ] <- NA

},spots$ID, n.samps, spots[,!(names(spots) %in% names(uniq.chip))])

}

aves <- data.frame(aves)

names(aves) <- names(spots)[!(names(spots) %in% names(uniq.chip))]

aves <- cbind(uniq.chip, aves)

write.table(aves, outfile.ave, sep="\t", row.names=FALSE)

print(proc.time())

q(save="no")

6. Generate chromosome plots using normalized data—pdf files can be viewed with Acrobat Reader.

## This program reads in the normalized BAC-averaged data and creates a

## chromosome-by-chromosome plot for each tumor.

## A postscript file is created for *each* array

norm.ave.file <- "Samples-20040621-norm-ave.txt"

centromere.file <- "chrom_info_GPApr2003.txt"

## Do you want to provide names for the program to use when labelling

## the sample plots? (TRUE/FALSE) If 'use.my.names' is FALSE then

## the column names from the file 'norm.ave.file' will be used.

use.my.names <- FALSE

my.samp.names <- c(

)

## read in data

aves <- read.table(norm.ave.file, sep="\t", header=T, as.is=T)

## drop BACs having no position information (if not previously removed)

aves <- aves[!is.na(aves$Mid),]

## Rescale "Mid" values so they are in Mbp rather than bp

aves$Mid <- aves$Mid/10^6

## get sample names

samp.names <- names(aves)[!(names(aves) %in% c("ID","Chrom","Mid","Start","End"))]

uniq.chrom <- sort(unique(aves$Chrom))

## read in centromere data

cent <- read.table(centromere.file, sep="\t", header=TRUE, as.is=TRUE)

options(papersize='letter')

for (i in 1:length(samp.names)) {

pdf(paste(samp.names[i],".pdf",sep=""),horizontal=FALSE,width=7.5,height=10)

#postscript(paste(samp.names[i],".ps",sep=""),horizontal=FALSE)

par(mfrow=c(4,2),oma=c(0,0,1.5,0), mgp=c(2,.5,0), mar=c(4,3,3,1.5)+.1)

for (j in uniq.chrom) {

ind <- (aves$Chrom==j)

y.lim <- c(-1,1)

y.lim <- range(c(y.lim,aves[ind,samp.names[i]]),na.rm=TRUE)

plot(aves$Mid[ind], aves[ind,samp.names[i]],

xlab="Distance along Chromosome (in Mbp)",ylab="Normalized Log-Ratio",

xlim=range(c(0,aves$Mid[ind]),na.rm=TRUE),

ylim=y.lim, main=paste("Chromosome",j), pch=20,

cex=.5, cex.lab=1.2, cex.main=1.35)

# cex=.5, cex.lab=1.2, cex.main=1.35)

abline(h=0)

## put line segment on plot corresponding to centromere

pos <- cent$cent.pos[cent$Chrom==j]/10^6

segments(pos,y.lim[1],pos,y.lim[2],lwd=1.5)

if (which(uniq.chrom==j) %in% seq(1,23,8))

mtext(paste("Sample: ",samp.names[i],sep=""),outer=T,cex=1.3)

}

dev.off()

}

q(save="no")

Quality control evaluation of array data

Section 12 Analysis of DNA for Genetic Instability and General DNA protocols

Multiple Displacement Amplification (MDA) using GenomiPhi kit

Amersham #25-6600-01

Protocol for sorted cells

1. Sort cells or nuclei into PCR tubes containing 10 ml mineral oil. Store tubes in the freezer at –20^oC until use.

2. Thaw cells on ice. Spin briefly to pellet cells. Estimate volume of cells/nuclei. (for 400 cells, ~1 ml after sorting)

3. Ideally, can add 1 ml of cell lysate to each 20 ml GenomiPhi reaction so adjust volumes accordingly. (*If volume is <0.1 ml, assume volume is zero)

4. Use either alkali or proteinase K lysis. For alkali lysis, mix an equal volume (or 0.5 ml) of lysis buffer: 400 mM KOH, 100 mM DTT, 1 mM EDTA. Incubate at RT for 10 minutes. Add an equal volume (or 0.5 ml) of neutralization buffer (400 mM HCL, 600 mM Tris-Cl). Mix gently.

5. For proteinase K, add volume of buffer: 1X TE, 1% Tween 20, 0.4 mg/ml proteinase K. Incubate samples at 55^oC for 3 hours.

6. Transfer 1 ml of the crude lysate to a new tube and mix in 9 ml GenomiPhi sample buffer.

7. In a separate tube,. mix 1ml GenomiPhi enzyme with 9 ml GenomiPhi reaction buffer per sample. Add 10 ml of this enzyme-reaction buffer mix to each sample tube.

8. Incubate samples at 30^oC for 3 hours.

9. Heat inactivate enzyme at 65^oC for 10 minutes.

10. Remove aliquot for DNA quantiation by Pico Green and store remainder of reactions at –20^oC.
*Expected yield of ~1-2 mg after 3 hours when starting with <1ng DNA

DNA purification using QIAGEN QIAEXII kit

*Use this kit to purify DNA fragments ranging in size from 40 bp-50 kb.

1. Thaw samples on ice. Verify sample volume.

2. Mix 3X volume QX1 buffer and 2X volume water.

3. Vortex QIAEXII beads for 30 seconds to resuspend.

4. Add 10 ml beads to each tube. Incubate for 10 minutes at RT, flicking gently every 2 minutes to keep beads resuspended.

5. Spin tubes at 12000 rpm for 20 seconds. Discard supernatant.

6. Wash pellet with 500 ml l PE buffer twice. (Do not resuspend pellet—just let buffer stand for 3-5 minutes.)

7. Spin tubes at 12000 rpm for 30 seconds.

8. Remove ALL remaining supernatant.

9. Air dry pellet for 15 minutes or until pellet turns white.

10. Add 15-20 ml TE per tube and flick gently to resuspend pellet. (Do not vortex—this could shear the DNA.)

11. Incubate tubes at 50^oC for 10 minutes to elute the DNA.

12. Spin tubes for 30 seconds. Transfer supernatant to new tube.

Re-quantitate DNA yield using Pico Green.

Arbitrarily Primed PCR (AP-PCR)

AP-PCR detailed protocol revised 4/26/04 by Lisa Lai

Methods are the same as Nancy Linford’s protocol except for the following revisions (see below). Following sample preparation, perform analysis as outlined previously.

PCR reactions (adapted from Rosana’s protocol—primer labeling step is omitted)

· Make the reagents up in order as a cocktail and add to DNA in 0.2 ml tube. Keep reagents on ice and ensure that they have thawed completely - vortex to mix

· Once the primer is added, all tips must go in acrylic box and reactions must be behind shield.

Sterile Water	6.3 ml	Hospital grade
10xPCR buffer with MgCl₂	1 ml
Primer (100 ng/ml)*	0.1 ml	MCG1 primer
MgCl² (25 mM)	0.4 ml
dNTPs (10 mM) *	0.1 ml
³²P-dCTP	0.1 ml	(1mCi/rxn; can decrease to 0.5mCi)
Taq polymerase	0.5 ml	Promega
DNA (5 ng/ml)**	1 ml

Primer: MCG1 5’-AAC CCT CAC CCT AAC CCC AA-3’
*aliquot to reduce freeze-thaw cycles

**measure DNA concentration with Pico Green and dilute accordingly

· Amplify DNA using the “APPCR” program on Thermocycler

AP-PCR Program:

94°C 3 min

Low stringency portion (5 cycles)

94°C 30 sec

45°C 1 min

72°C 1.5 min

High stringency portion (35 cycles)

94°C 15 sec

50°C 15 sec

72°C 1 min

72°C 7 min

4°C hold

Store samples at –20^oC after amplification is completed

Gentra+ Puregene DNA Isolation from Sorted Nuclei

Cell Lysis

1. Go to room K094 (at 4° C).

2. Make sure samples are thawed on ice before putting them in centrifuge. Centrifuge sorted nuclei at 14,000rpm on Eppendorf Centrifuge 5415D (13,000-x g, 6cm radius) for 2 minutes to pellet cells.

3. Working on ice, pipette off supernatant or drain on Sorbeez pad (if volume >500 ml) leaving <100 ml liquid. 100ul is approximate. 50ul-150ul should be fine. You can also pipette off greater than 500ul if you like.

4. Thaw proteinase K (20 mg/ml) in ice bucket. Add 200 ml Cell Lysis Solution and 10 ml proteinase K to each tube, and Vortex. Keep on ice until cell lysis is added (you can drop from above the 200ul using only one pipette tip if you don’t put it in the tube). You can add proteinase K off ice and then at this stage they can be off ice indefinitely.

5. Mix by flicking the tube. Place tube in 50°C water bath overnight. After mixing you should see bubbles/ foam. If not cell lysis was forgotten.

RNase Treatment

6. Flash spin.

7. Add 1.0 ml RNase A Solution to the cell lysate.

8. Mix the sample by inverting the tube 25 times.

9. Incubate at 37° C for 15-60 minutes. No difference in cell recovery based on time at this stage.

Protein Precipitation

10. Flash spin. Cool the sample to room temperature.

11. Add 66 ml Protein Precipitation Solution. You can drop from above the 66ul with one pipette tip if you don’t touch the tube. This can be done off ice.

12. Vortex for 20 seconds.

13. Centrifuge at 10,500 rpm (10,000g, 8cm radius) in Eppendorf Centrifuge 5417C for 10 minutes at room temperature. The precipitated proteins will form a tight pellet.

DNA Precipitation

14. Add 200 ml Isopropanol to a new 1.5-ml tube (add 1ul of glycogen if cells numbers are lower than 200k). Your new tubes need to be labeled on side with name, flow, level and top with DNA number, fraction (G1/4N, etc) and cell count before transferring original DNA sample.

15. Pipette off the supernatant containing the DNA (leaving behind the precipitated protein pellet) and add to the Isopropanol in the new 1.5-ml tube. You can put your tip down to meniscus and draw up until slightly over the visible protein pellet. Usually this will take 2 pipettes. I use 200ul tips. Don’t use larger b/c tip is too big and will suck up protein pellet. If protein pellet is not visible something is wrong. Start at step 11 again if you don’t see a pellet. If you dislodge pellet or accidentally suck up part of the protein pellet you need to start at step 13 again because it will mix with the DNA. Normal tends to be very thick at this stage (almost mucous like) and proceed carefully b/c it’s easy to dislodge the pellet w/ normal.

16. Mix the sample by inverting gently 50 times. Allow to precipitate at -20°C overnight. You can precipitate for indefinite amount of time (nights, week, etc)

17. Centrifuge at 11,500 rpm (9000g, 6cm radius) in Eppendorf Centrifuge 5415D in Room K094 for 30 minutes at 0°C.

18. Pour off the supernatant and drain tubes on a clean absorbent paper. There will be no visible pellet at this stage.

19. Add 500 ml 70% EtOH and invert the tubes several times to wash the DNA pellet. Doesn’t need to be on ice at this stage and below.

20. Centrifuge at 10,500 rpm (8000g, 6cm radius) in Eppendorf Centrifuge 5415D in Room K094 for 10 minutes. Carefully pour off the ethanol.

21. Repeat steps 20 and 21.

22. Allow to air dry overnight or speed-vac on medium heat until dry (around 30 minutes). Air dry unless in a big hurry. The air dry can be several nights (indefinite) if needed.

DNA Hydration

23. Add 1 ml DNA Hydration Solution for every 1,000 cells. For example, add 10 ml DNA Hydration Solution to 10,000 sorted cells. If you’ve only dried over one night check each tube for remaining liquid. If found heat vacuumed dry or wait to rehydrate sample.

24. Heat at 65°C for 1 hour. Tap tube periodically to aid in dispersing the DNA.

Record of Reagents Used

Cell Lysis Solution:

20 mg/ml proteinase K:

RNase A:

Protein Precipitation Solution:

Isopropanol:

Glycogen:

70% EtOH:

DNA Hydration Solution:

Last updated 07/28/2003, PG & LJ

Further revised 07/31/2003, KL

PICOGREEN PROTOCOL TO QUANTIFY DNA IN FLUOROCOUNTER

Set up samples

1. Prepare a standard curve with a reference DNA (for example, Lambda DNA from Invitrogen).

2. Thaw the picogreen dye at 37C (DMSO freeze point is 18.55°C)

3. Dilute picogreen in TE (1/400). For one plate: 10ml TE and 25ul picogreen.

4. Mix the dye solution and add 95ul to each well of the black 96-well plate.

5. Add 5ul of DNA (fluorocounter saturates at 50ng, dilute accordingly) to each well.

6. Add 5ul of standard curve and. It’s better not to use any positive control for the max RFU position. Use just a standard curve. See Fluorocount Manual Pg 84 or below.

7. Let the mix sit in the dark for 5-10 min.

8. Test the samples on the fluorocounter.

Fluorocounter

1. Turn the fluorocounter and the light-source on (for maximum efficiency the light-source should be on 15min before analyzing. Leave a note on the machine indicating that is on use).

2. Sign into the fluorocounter log book.

3. Double-click fluorocount icon

4. Click ‘Plate out’ icon

5. Load your plate into the machine and click the ‘Plate in’ icon

6. Click File, Open, rosana.pf, OK

7. Click Plate, Read settings, Read modeTab. Type Upper Left position (i.e. A01) and Lower Right position (i.e. H12). The rest is by default.

8. Click Sensitivity Tab. Leave max RFU blank and select read length 2.0, gain 1.0 and PMT 1100. These are the optimal settings. If a RFU well is indicated then the program doesn’t use the optimal settings but the ones appropriate for the max RFU. Since the max RFU changes between plates, this introduces variability in the comparison of data of different plates. For concentrations of DNA between 2 and 0.06 ng/ul (10ng and 0.3ng) use gain 7.0 PMT 1100.

9. Click Filter pairs Tab. Scan 2. Ex: 485, Em: 530

10. Click OK to exit.

11. Click Read Plate icon (looks like a large P). You should see the UL well, LR well and the Max RFU well that you selected in step 6 and 7). Click Read (it should produce numbers in all of the wells you selected). Gain and PMT are autoset: the fluorocounter selects the best values to read the plate. That’s why if RFU is too concentrated the sensitivity to read the rest of plate will be lower.

12. Close the window when it is finished by clicking cancel.

13. Click File/export. Export to select Microsoft Excel. Change the name of the file to what you want it to be. Click on Export Dir to select your folder in SpectraCount. Click on the Data to Export tab. Scroll down to the last reading and select it (it should be something like {110, 1/1}). Click Export. It should show you the name that you selected. Click Export. It will say Export is complete when done. Click OK to close the box.

14. Remember to take out your plate and if you are done then exit the program.

15. You can e-mail the file to yourself, it’s a few Kb.

16. Close the windows, shut off the light source and the flourocounter. If you plan to use it in a hour or so you can keep the light source on. Leave a note indicating that you’d come back.

NOTE: The fluorocounter has a program called i-smart to analyze the plate. You can use it or make your own calculations in Excel.

Black plates: 300ul, black polypropylene, flat bottom, Whatman 7701-7350 (101.20$/50plates). The plates can be washed and recycled. They can be autoclaved too.

PicoGreen® dsDNA quantitation reagent *200-2000 assays*, 1 mL. Molecular Probes P7581 www.probes.com

Gel Electrophoresis Protocol

Preparation of 0.7% agarose

- Add 1.75g agarose and 250ml of 1X TAE (tris-acetate) to a 500ml Erlenmeyer flask

- Heat mixture in microwave until agarose is totally dissolved

- Cool mixture with tap water

- Add 12.5ul EtBr (10mg/ml)

- Continue cooling mixture to about 40-50 degrees Celsius

- Pour into long gel rig

- Wait for 30 minutes

Preparation of 1% agarose

- Add 2.5g agarose and 250ml of 1X TBE (tris-borate) to a 500ml Erlenmeyer flask

- Heat mixture in microwave until agarose is totally dissolved

- Cool mixture with tap water

- Add 12.5ul EtBr (10mg/ml)

- Continue cooling mixture to about 40-50 degrees Celsius

- Pour into long gel rig

- Wait for 30 minutes

Preparation of Buffer Solution (replace TAE with 1X TBE if TBE is used)

- Mix 2L 1X TAE (40ml 50X TAE + 1960ml water) and 0.5ug/ml EtBr (100ul 1% EtBr solution)

- Swirl

Loading Samples and Markers

- Spin down all samples

- Get a plastic plate (353911)

- For samples: Add 2ul of 6X LB (loading buffer; obtained in 4 degrees fridge) and 10ul of sample to each well on the plate

- For markers: Add 4ul of 100bp DNA ladder (obtained in 4 degrees fridge)

- Load the gel, using unfiltered pipet tips

- Plug wires into power source

- Set at approximately 120V

- Run the gel for 2 to 6 hrs

To view the gel

- Disconnect from power source

- Wash gel with distilled water

- Go to UV room and shine UV light on gel (turn lights off)

- Zoom in/zoom out and adjust contrast

- Print picture of gel

Note: All amounts can be scaled depending on the amount of gel needed and the DNA to be analyzed.

Last updated by KL 4/30/04

Southern Blotting Protocol

Depurination

- Soak gel in 0.25M HCl for 15 minutes in glass tray

- Discard HCl

Denaturation

- Soak gel in at least 2 gel volumes (~500ml) of 1.5M NaCl/0.5N NaOH for 30 minutes in glass tray

- Discard liquid

Set up

- Stack brown paper towels to height of 5cm

- Cut 4 pieces of cardboard paper (obtained in drawer) to the same size as gel

- Put 2 cardboard paper on top of paper towels

- Cut nylon membrane to the same size as gel

- Put membrane on top of cardboard paper

- Put gel (upside down) on top of membrane

- Put 2 cardboard paper on top of gel

- Soak 2 sponges in buffer solution and put them on top of cardboard paper

- Surround the gel with plastic wrap

- Wrap the whole thing with plastic wrap

Preparation of Pre-hyb. Solution

- Add 10ml water to tube using pipet

- Add 6ml 20X SSC (store in lab)

- Add 2ml 10% SDS (prepare 1L pH 7.2)

- Add 2ml 50X Denhardt’s (melt in boiling water)

- Add 66ul 10mg/ml Herring Sperm DNA

Preparation of Hyb. Solution

- Add 10ml water to tube using pipet

- Add 6ml 20X SSC

- Add 2ml 10% SDS

- Add 2ml 50X Denhardt’s (melt in boiling water)

- Obtain 50ul of Probe (7.6*10^6 cpm)

- Denature 5’ in boiling water

- Put in ice

- Add Probe to Hyb. solution using a pipet

Hybridization

- Cut mesh as the size of the membrane

- Coil the mesh and membrane and put in bottle

- Prewet in 2X SSC and warm in 37 degrees

- Prewarm Pre-hyb. and Hyb. solution at 37 degrees

- Discard 2X SSC and add Pre-hyb. solution

- Warm at 37 degrees for 2 hours

- Discard Pre-hyb. and add Hyb. solution

- Warm at 37 degrees overnight

Washes

- Discard Hyb. solution and add 50ml 2X SSC

- Warm at 37 degrees for 15 minutes

- Discard SSC and add 50ml 2X SSC

- Warm at 37 degrees for 15 minutes

- Discard SSC and add 50ml of 2X SSC and 0.1% SDS

- Put at room temperature for 30 minutes

- Discard solution and add 50ml of 0.1X SSC

- Put at room temperature for 5 minutes

- Take out membrane

- Dry membrane

- Blot membrane on Whatman paper

- Wrap membrane in plastic wrap

- Expose to phosphorimage screen for 1.5 hours

Last updated by KL 6/30/03

Phenol Chloroform extraction for Buffy Coat Cells or other tissue

Protocol modified from Jon E Digel at FHCRC (11-4-03)

Notes and Precautions

All buffy coat samples should be handled in accordance with procedures specified in the Blood Bourn Pathogen course (gloves, lab coat, and face protection). Autoclave all plastics before disposal.

Principle

Buffy Coats from prepared blood samples, drawn into vacutainer tubes containing EDTA as anticoagulant for epidemiological studies, are stored at -80^oC in a media that maintains the integrity of the cells under freezing conditions. The buffy coat is washed twice prior to DNA extraction. [Washing removes the components of any freezing media as well as any contaminating red blood cells that may interfere with PCR and other molecular biology procedures.] The washed white cells undergo digestion in a solution of 10% SDS and the enzyme Proteinase K (Protease). What remains is the DNA that has been released from the cell nucleus. Two phenol:chloroform extractions are performed to remove the digested proteins. A single chloroform extraction is performed to remove any residual phenol. DNA is then precipitated out of the aqueous solution using 100% ethyl alcohol and a salt solution of sodium acetate. [More than one precipitation with cold 100% ETOH may be performed at cold temperatures (-20°C) to deplete a sample of all DNA.] The DNA precipitate is then washed with 70% ethyl alcohol to remove any remaining salts. Pure DNA is then dissolved in buffer and quantitation (mg/mL) determined by spectrophotometry. A final concentration (ng/uL) is produced based on the needs of the recipient.

Equipment and Supplies

Equipment

repeating pipetter, adjustable from 1 mL - 50 mL

rocker platform

refrigerated centrifuge

water bath temperatures up to 70^oC

vortex mixer, such as Thermolyne

spectrophotometer capable of reading 260/280 wavelengths

Supplies

centrifuge tubes, sterile, 15 ml, polypropylene, conical bottoms with caps

Eppendorf, microcentrifuge conical tubes

syringe-style pipet tips, 50 mL capacity

disposable transfer pipettes

beakers of various sizes

reagent bottles, 500 mL capacity

styrofoam racks for centrifuge tubes

UV transparent disposable plates, 96 wells

Chemicals

1X PBS, sterile Made in lab

Proteinase K, 500 mg Roche

Tris pH 7.5, or pH 8.0 Made in lab

NaCl Fisher Scientific

EDTA, pH 7.5 or 8.0 Made in lab

10 % sodium dodecyl (Lauryl)sulfate in water (SDS) prepared in lab

ETOH 200 proof (100%) Fisher Scientific

3M sodium acetate, pH 5.0 prepared in lab

Phenol - buffer saturated, pH 6.7 +/- 0.2 Fisher Scientific

CAUTION: Organic Solvent! Goggles and gloves must be worn when handling this chemical

Chloroform - purified, molecular biology grade Fisher Scientific

CAUTION: Organic Solvent! Goggles and gloves must be worn when handling this chemical

Laboratory’s preparation of WORKING solutions:

24:1 phenol/chloroform mixture To extract 48 samples:

Add 144 mL phenol

6 mL chloroform

1X STE

1 liter stock solution: 20X STE Add 24.23 g Tris

116.88 g NaCl

7.45 g EDTA

QS to 1000 mL with ddH₂0, pH to 8.0

Autoclave or filter sterilize

Store at room temperature

Indefinite expiration

Laboratory prepares 500 mL working solution: 1X STE 25 mL stock

475 mL ddH₂0

Store at room temperature

Indefinite expiration

10% SDS.

1 liter working solution: 10% SDS Add 100 g SDS to 1000 mL ddH₂0

Store at room temperature

Indefinite expiration

3M NaOac (sodium acetate)

500 mL working solution: 3M Add 123.1g NaC₂H₃O₂ to 500 mL ddH₂O,
pH to 5.2

Autoclave or filter sterilize

Store at room temperature

Indefinite expiration

Proteinase K (Freezer 4 Rack 9 Box 1)

Laboratory prepares 10 mL working solution: 50 mg/mL Add 10 mL ddH₂0

to one vial of 500mg Proteinase K

Store at -20^o C

Indefinite expiration

70% ETOH

1 liter working solution: 70% ETOH Add 700 mL of 100% ETOH

300 mL ddH₂0

Mix well

Store at 2-8°C

Indefinite expiration

TE Buffer

1 liter stock solution: 20X TE buffer Add 24.23 g Tris

7.45 g EDTA

QS to 1000 mL with ddH₂0, pH to 7.5

Autoclave or filter sterilize

Store at room temperature

Indefinite expiration

500 mL working solution: 1X TE buffer Add 25 mL stock TE buffer

to 475 mL ddH₂0

Store at room temperature

Indefinite expiration

Procedure

Sample Preparation and DIGESTION (Day 1):

Samples are to be thawed in the refrigerator.

Transfer the thawed samples into appropriately labeled, sterile 15 mL centrifuge tubes. Add 5 mL 1X PBS. Mix well.

Centrifuge for 15 minutes 1500 rpm, at RmT°.

Pour off the supernatant. Samples are to be visually inspected for the presence of red blood cell/hemolysis contamination for reference during protein digestion and measurement (mm) of the buffy coat.

To the pellet, add 6 mL of 1X STE. Manually agitate the conical tubes to break up cell pellets.

Add 150 mL of 10% SDS and 50 mL of working solution Proteinase K to digest cells. Mix well.

Incubate overnight at 55°C in a water bath. Gently mix several times over the course of the first hour
or two.

EXTRACTION and PRECIPITATION PROCEDURE (Day 2):

Add 5 mL phenol (an equal volume to the volume in the conical tube) to the 15 mL conical tube.

Using a rocker, rock the samples at room temperature for 20 minutes.

Centrifuge for 20 minutes at 3000 rpm, RmT°.

Transfer the supernatant (the upper layer) to a clean 15 mL conical tube.
NOTE: The phenol is disposed of in phenol waste in the fume hood.

Add 5 mL chloroform (an equal volume to the volume in the conical tube) to the 15 mL conical tube.

Using a rocker, rock the samples at room temperature for 20 minutes.

Centrifuge for 20 minutes at 3000 rpm, RmT°.

Add two volumes (approximately 12 mL) of 100% (200 proof) ETOH and 600 mL (or 1:10 volume of the supernatant) of 3M NaOac, pH 5.2. Place on ice for 10 minutes.

Shake vigorously for 2 minutes, or rock for 1 hour, to precipitate all of the DNA.
[Notice the flocculation of the DNA as the conical tube is first agitated.]

NOTE: Since small volumes of DNA are expected, the tubes may be held overnight at -20°C to facilitate precipitation of all of the DNA present. Allowing the tubes to remain at low temperatures will INCREASE the salt concentrations.

Centrifuge for 30 minutes at 3500 rpm, RmT°.

Remove and discard the ETOH/NaOac by aspiration or decant.

Add 5 ml 70% ETOH to the conical tube and gently dislodge the pellet from the bottom of the tube. DO NOT shake. [Notice the crystalline presence (salt) at the interface of the pellet and the dilute alcohol. Rock for 1 hour at RT.This step dissolves the salts that precipitated out of solution along with the DNA.]

Centrifuge for 20 minutes at 3500 rpm, RmT°.

Remove and discard the 70% ETOH by aspiration or decant.

Air dry the pellet for 30 minutes. DO NOT over dry the pellet.

Add 1 mL 1X TE buffer to the conical tube dependent upon amount of DNA present) and again dislodge the pellet from the bottom of the tube. Allow the pellet to dissolve overnight at 38+/- 2°C in a shaking incubator.

QUANTITATION PROCEDURE (Day 3):

Read the absorbance at 260/280 on the Smartspec. A 1:50 dilution is good. Save the DNA concentration print out in mg DNA/mL. Aliquote 100 ul into a 0.5 ul tube for Rosana and put in the refrigerator. Put the remaining DNA into a 1.7 ul tube and freeze.

Limitations

Pure DNA has a A260/280 ratio of 1.8. An allowable performance limit for each specimen is a A260/280 ratio of > 1.70 and < 2.14. If a ratio is outside of the allowable limit, the specimen must be re-processed. Ratios below 1.75 are indicative of proteins being present in the final solution. Ratios greater than 2.10 are indicative of salts being present in the final solution. If the presence of proteins is suspected, the specimen is to be re-extracted. If the presence of salts is suspected, the specimen is to be re-precipitated using 100% ethanol and washed overnight with 70% ethanol.

Low concentrations of DNA with good A260/280 ratios are acceptable for PCR.

High concentrations of DNA with poor A260/280 ratios need to be allowed to dissolve longer and then reread.

The linearity of the indicates that L1 values (OD260) below 0.01 and above 2.5 do not provide dependable values. Any reading outside of this range MUST be repeated at an appropriate dilution.

References

Innis M.A., Gelfand D.H., Sninsky J.J., White T.J., PCR Protocols: A Guide to Methods and Applications, Harcourt Brace Jovanovich, publ., 1990, p.148.

Farkas D.H., Molecular Biology and Pathology: A Guidebook for Quality Control, Harcourt Brace Jovanovich, publ., 1993.

McDougall J., McDougall HLA Lab Standard Procedures.

Maniatis T., Fritsch E.F., Sambrook J., Molecular Cloning, A Laboratory Manual, Cold Spring Harbor, publ, 1989.

Protocol: QIAamp Blood Kit, Rapid Purification of DNA for Direct Use in PCR Amplification, QIAGEN Inc. 9600 De Soto Avenue, Chatsworth CA 91311.

Edwards A.M., Hunter S.V., Hankin R.C., Gene Rearrangement Analysis by Nonorganic Extraction and Chemiluminescent Detection, Laboratory Medicine: 24 (10), Oct 1993.

SOFTmax PRO Protocol Guide, Version P1.12, Molecular Devices Corporation, Sunnyvale, CA 94925.

Section 13 Solutions

AMP

To make 25 mg/ml AMP:

2.5 g ampicillin

100 ml d H₂0

1. Swirl well (don’t need stir bar).

2. Filter-sterilize.

3. Aliquot into 25 - 5ml snap-cap tubes (4 ml per tube).

4. Label “25 mg/ml AMP”.

5. Store in the freezer.

Reagents Information:

Ampicillin: Location – Refrigerator #1, Shelf #3, Box #5

Manufacturer – Shelton Scientific; Cat# IB02040

AMP PLATES

To make 1 L:

20 g LB broth

10 ml 1M MgSO₄

10 g Bacto-Difco agar

qs d H₂O to 1 L

Notes: Make this solution in two 500ml batches. Bacto-Difco agar doesn’t dissolve so divide it into each 500 ml solution when making up 1 L.

For LB plates, don’t add AMP.

1. Autoclave then cool in 55^˚C bath

2. Add 2 ml of 25 mg/ml AMP to 500 ml LB AGAR solution.

3. Pour 15 ml into each plate.

Reagents Information:

LB broth: Location – Lower Shelf 1

Manufacturer – DIFCO Industries; Cat# 0446-17-3

MgSO₄: Location – Upper Shelf 1

Manufacturer – Sigma; Cat# M-1880

Bacto-Difco agar: Location – Lower Shelf 1

Manufacturer – Becton Dickinson; Cat# 214010

AGAROSE for DNA gels

Reagents Information:

Sea-Kem agarose: Location – Lower Shelf 1

Manufacturer – FMC BioProducts; Cat# 50001

TBE (10X): Location – Liquids Shelf (brown metal shelf unit third bay, left side)

Manufacturer – Bio-Rad; Cat# 161-0770

TAE (50X): Location – Middle Bay, Left side, Shelf 1

Manufacturer – Bio-Rad; Cat# 161-0743

1st BLOT WASH (2X SSC, 0.1% SDS)

** WEAR MASK WHEN WORKING WITH SDS! **

To make 4 L:

4 g SDS

400 ml 20X SSC

qs dH₂O to 4 L

Dissolve SDS in 1 L dH₂O. Then add SSC and qs dH₂O to 4L.

Reagents Information:

SDS: Location – Upper Shelf 1

Manufacturer – BDH; Cat# 442444H

SSC: Location – Liquids Shelf (brown metal shelf unit third bay, left side)

Manufacturer – Bio-Rad; Cat# 161-0775

2nd BLOT WASH

** WEAR MASK WHEN WORKING WITH SDS! **

To make 4 L:

4 g SDS

20 ml 20X SSC

qs dH₂O to 4 L

Dissolve SDS in 1L dH₂O. Then add SSC and qs dH₂O to 4L.

Reagents Information:

SDS: Location – Upper Shelf 1

Manufacturer – BDH; Cat# 442444H

SSC: Location – Liquids Shelf

Manufacturer – Bio-Rad; Cat# 161-0775

Calcium Chloride Sheath Fluid (5mM)

Use for sorting flow cytometry samples onto slides

To make 6 L of 5mM:

6 L dd H₂O

4.41 g CaCl₂

1. Pour 5 L dd H₂O into clean 6-L flask.

2. Weigh out the CaCl₂.

3. Using powder funnel, add solid calcium chloride to flask while stirring.

4. Add the last liter of dd H₂O; try to rinse the inside of weigh boat, powder funnel, and flask with this remainder dd H₂O to bring the total ddH₂O to 6 liters.

5. Stir until calcium chloride is dissolved.

6. Get new bell filter and tubing from drawer labeled "CaCl₂ tubing".

7. Filter the solution: Use the peristaltic pump; switch PBS tube (with filter attached) to the CaCl₂ tube. Run pump for a few seconds without the new bell filter attached to rinse the tube.

8. Put new bell filter on end of tube.

9. Pump CaCl₂ solution (from the flask the solution was mixed in) into CaCl₂ bottle.

10. Return CaCl₂ bottle to walk in refrigerator.

Reagents Information:

CaCl₂: Location – Lower Shelf 1

Manufacturer – Sigma; Cat# C-3881

1X DAPI (Working Solution)

To make 1 L:

500 ml ddH₂0

8.5 g NaCl (final conc. = 146 mM)

1.2 g Tris Base (final conc. = 10 mM)

1. Add these 1^stthree ingredients together.

2. Adjust pH to 7.4 with HCl.

3. Add:

a. 4 ml of 500 mM CaCl₂solution (final conc. = 2 mM)

b. 44 ml of 500 mM MgCl₂solution (final conc. = 22 mM)

c. 50 mg (0.05g) BSA

d. 1 ml Nonidet P-40 detergent (aka: Igepal) (final conc. = 0.1%)

e. 10 mg DAPI (4,6-diamidino-2-phenylindole) powder (final conc. = 10 ug/ml)

f. 100 ml DMSO (final conc. = 10%)

4. Add dd H₂0 to final volume of 1 L.

5. Store in dark or foil wrapped bottle at 2-6^˚C.

Reagents Information:

NaCl: Location – Upper Shelf 1

Manufacturer – Fisher Scientific; Cat# S271-3

Tris Base: Location – Upper Shelf 1

Manufacturer – Fisher Scientific; Cat# BP152-1

CaCl₂: Location – Lower Shelf 1

Manufacturer – Sigma; Cat# C-3881

MgCl₂: Location – Upper Shelf 1

Manufacturer – Sigma; Cat# M-8266

BSA: Location – Refrigerator #1, Shelf #2, Box #4

Manufacturer – Sigma; Cat# A-6003

Nonidet P-40: Location – Upper Shelf 1

Manufacturer – Sigma; (Igepal CA-630)Cat# I-3021

DAPI: Location – Refrigerator #1, Shelf 2, Box #3

Manufacturer – Accurate Chem. Co.; Cat# 18860 (No Substitutes)

DMSO: Location – Lower Shelf 1

Manufacturer – Fisher Scientific; Cat# D128-500

DENATURING SOLN for Southerns

To make 2 L:

2 L water

175.32 g NaCl

40 g NaOH

Reagents Information:

NaCl: Location – Upper Shelf 1

Manufacturer – Fisher Scientific; Cat# S271-3

NaOH: Location – Upper Shelf 1

                                Manufacturer – Fisher Scientific; Cat# S318-500

DEPC TREATED WATER

** CARCINOGEN. WORK UNDER HOOD! **

500 ml dH₂O

8 drops DEPC (diethyl pyrocarbonate, stored @ 4°C)

1. Cap bottles and shake until foams.

2. Let sit loosely capped @ RT for 1 hr+ (to remove RNase from dH₂0).

3. Autoclave to degrade DEPC to alcohol and CO₂.

Reagents Information:

DEPC: Location – Refrigerator #1, Shelf #1, Box #1

Manufacturer – Sigma; Cat# D-5758

0.5M EDTA pH 8.0

To make 1 L:

186.1 g EDTA

Add to 800 mls dH₂O

1. pH has to be over 8 for it to go into solution.

2. Add NaOH PELLETS to raise pH.

3. Add pellets until solution clears.

4. qs dH₂O to 1 L.

Reagents Information:

EDTA: Location – Lower Shelf 1

Manufacturer – JT Baker, Inc.; Cat# 4040-01

NaOH: Location – Upper Shelf 1

Manufacturer – Fisher Scientific; Cat# S318-500

Hoechst 33342 (viable DNA stain) 1mM

To make 200 ml:

0.112 g Hoechst 33342 (carcinogenic handle with gloves, mask and lab coat)

200 ml of dH₂0

2 ml 95% ethanol

1. Mix thoroughly.

2. Cover the bottle with aluminum foil to protect from the light.

3. Store at 4°C.

Reagents Information:

Hoechst 33342: Location – Freezer #1, Box #1

Manufacturer – Sigma; (Bisbenzamide Hoechst 33342) Cat# B-2261

95% ethanol: Location – Flammables Cabinet

Manufacturer – Aaper Alcohol; Cat# 03E22QA

LB BROTH

To make 1 L:

20 g LB broth base

10 mls 1M MgSO₄

qs dH₂ O to 1 L then autoclave

Reagents Information:

LB broth: Location – Lower Shelf 1

Manufacturer – DIFCO Industries; Cat# 0446-17-3

MgSO₄ : Location – Upper Shelf 1

Manufacturer – Sigma; Cat# M-1880

DNA Loading Dye for Acrylamide Gel

To make 10 ml:

95% Deionized Formamide 9.5 ml Deionized Formamide

0.1% Bromophenol Blue 1.0 mg Bromophenol Blue

0.1% Xylene Cyanol 1.0 mg Xylene Cyanol

10 mM EDTA 2.92 mg EDTA (FW = 292.2 g)

0.5ml dH₂O

Use in a 1:1 ratio to the DNA sample.

Reagents Information:

Deionized Formamide: Location – Refrigerator #1, Main shelf #3,

Manufacturer – Amresco; Cat# 0606-500ml

Bromophenol Blue: Location – Lower Shelf 1

Manufacturer – Sigma; Cat# B-7021

Xylene Cyanol: Location – Upper Shelf 1

Manufacturer – Sigma; Cat# X-4126

EDTA: Location – Lower Shelf 1

Manufacturer – JT Baker, Inc.; Cat# 4040-01

MitoTracker Green (MTG)

To make 200mM stock solution:

50 mg stock vial from Molecular Probes

372 ml DMSO

Store the 200mM stock solution in Freezer #2, Shelf #1, Box #1.

Reagents Information:

MTG (MW=671.88): Location – Freezer #4, Mitotracker Jar

Manufacturer – Molecular Probes; Cat# M-7514

DMSO: Location – Lower Shelf 1

Manufacturer – Fisher Scientific; Cat# D128-500

MitoTracker Red (CmxRos)

To make 200mM stock solution:

50 mg stock vial from Molecular Probes

470 ml DMSO

Store the 200mM stock solution in Freezer #2, Shelf #1, Box #1.

Reagents Information:

CmxRos (MW=531.52): Location – Freezer #4, Mitotracker Jar

Manufacturer – Molecular Probes; Cat# M-7512

DMSO: Location – Lower Shelf 1

Manufacturer – Fisher Scientific; Cat# D128-500

2% PARAFORMALDEHYDE

To make 500 mls:

10 g of paraformaldehyde

500 ml of 1x PBS (from TC) at pH of 12

1. Measure 500 mls of PBS into a 500 ml bottle and adjust the pH to 12.

2. Heat it to 60°C under a fume hood

3. Add 10 g of paraformaldehyde and stir for about 5 to 10 minutes

4. Cool until to room temperature under a fume hood

5. Finally adjust the pH to somewhere between 7 and 8

Reagents Information:

Paraformaldehyde:               Location – Upper Shelf 1

Manufacturer – Fisher Scientific; Cat# T353-500

PBS (10X): Location – Glassware Cabinets

Manufacturer – Invitrogen/Gibco BRL; Cat# 14200-075

10X PBS solution for Flow Cytometer

To make 5 L:

400 g of NaCl

10 g of KCl

10 g of KH₂PO₄

56.75 g of Na₂HPO₄

qs dd H₂O to 5 L

1. Pour ~ 4 L of dd H₂O into the 5-L flask marked with “10X PBS ONLY”.

2. Weigh out each solid compound listed above on the scale.

3. Using the powder funnel, add the solid compound into the flask while stirring.

4. When all four compounds have been added, add enough dd H₂O to fill the flask up to 5 L. Try to rinse the inside of the powder funnel and the inside of the flask near the mouth with the dd H₂O used to fill up to 5 L.

5. Continue to stir until all the compounds are dissolved in solution.

6. Then, using the liquid funnel, pour out 500ml into ten 500ml flasks marked with “10X PBS”.

Reagents Information:

NaCl: Location – Upper Shelf 1

Manufacturer – Fisher Scientific; Cat# S271-3

KCl: Location – Upper Shelf 1 (PBS Box)

Manufacturer – JT Baker; Cat# 7447-40-7

KH₂PO₄: Location – Upper Shelf 1 (PBS Box)

Manufacturer –JT Baker; Cat# 7558-79-4

Na₂HPO₄: Location – Upper Shelf 1 (PBS Box)

Manufacturer – JT Baker; Cat# 7778-77-0

1x PBS for Tissue Culture

To make 5 L:

500 ml bottle of Dulbecco’s PBS (10x)

4500 mls of bottled water

1. Mix and adjust pH to 7.1

2. Aliquot into 100 ml bottle, put caps on LOOSELY and tape with autoclave tape

3. Label with red tape (PBS, the date and your initials) and autoclave (in D wing only)

4. When bottles are completely cool, tighten caps.

Reagents Information:

Dulbecco’s PBS (10x): Location – Glasswares Cabinet

Manufacturer – Invitrogen/Gibco BRL; Cat# 14200-075

20% SDS

** WEAR MASK WHEN WORKING WITH SDS!! **

To make 500 mls:

100 g SDS

500 mls dH₂O

1. Add SDS to 400 mls dH₂O.

2. Stir.

3. Heat to 68^˚C to get into solution, then cool.
Then adjust the pH to 7.2 (very tricky- use dilute HCl/ NaOH).

4. Add dH₂O to 500 mls.

Reagents Information:

SDS: Location – Upper Shelf 1

Manufacturer – BDH; Cat# 442444H

50X TAE

To make 500 mls:

121.1 g Trizma base

50 ml 0.5M EDTA

28.55 ml glacial acetic acid (PROTECT EYES! )

qs dH₂O to 500 mls

Don’t need to autoclave.

Reagents Information:

Trizma base: Location – Upper Shelf 1

Manufacturer – Sigma; Cat# T-6066

EDTA: Location – Lower Shelf 1

Manufacturer – JT Baker, Inc.; Cat# 4040-01

Glacial Acetic Acid: Location – Acids Cabinet

Manufacturer – Fisher Scientific; Cat# A38S-500

TB (TERRIFIC BROTH):

To make 500 mls:

TB: 6 g bactotryptone

12 g bacto yeast

2 ml glycerol

dH₂O to 450 mls

KPO₄ (“K” buffer):

1.155 g KH₂PO₄

6.27 g K₂HPO₄

dH₂O to 50 mls

1. Autoclave the two solutions separately.

2. Add 50 mls of sterile KPO₄ buffer to 450 mls of TB before use.

Reagents Information:

Bactotryptone: Location – Lower Shelf 1

Manufacturer – DIFCO Industries; Cat# 0123-17-3

Bacto Yeast: Location – Lower Shelf 1

Manufacturer – Becton Dickinson; Cat# 212750

Glycerol: Location – Lower Shelf 1

Manufacturer –JT Baker; Cat# 2136-01 or Sigma; Cat# G-516

KH₂PO₄: Location – Upper Shelf 1

Manufacturer –JT Baker; Cat# 7558-79-4

K₂HPO₄: Location –Upper Shelf 1

                                                Manufacturer – JT Baker; Cat# 3252-01

10X TBE:

To make 1 L:

108 g Trizma base

40 ml 0.5 M EDTA

55 g boric acid

qs d H₂O to 1L

Don’t need to autoclave.

For gel use, dilute to 0.5 X TBE. (50 ml 10 X TBE and add dH₂O to 1000 ml.)

Reagents Information:

Trizma base: Location – Upper Shelf 1

Manufacturer – Sigma; Cat# T-6066

EDTA: Location – Lower Shelf 1

Manufacturer – JT Baker, Inc.; Cat# 4040-01

Boric Acid: Location – Lower Shelf 1

Manufacturer – EM Science; Cat# BX0870-1 or 10043-35-3

TE (10 mM TRIS, 1 mM EDTA)

To make 1 L:

10 ml 1M TRIS

2 ml 0.5 M EDTA

qs to 1 liter in dH₂O

Reagents Information:

TRIS: Location – Upper Shelf 1

Manufacturer – Fisher Scientific; Cat# BP152-1

EDTA: Location – Lower Shelf 1

Manufacturer – JT Baker, Inc.; Cat# 4040-01

1M TRIS

To make 1 L:

121.1 g Trizma base

800 ml dH₂O

add HCl to get to desired pH (WEAR GOGGLES! DO UNDER HOOD!)

qs dH₂O to 1L

Use glass pipettes when working with HCl.

Reagents Information:

Trizma base: Location – Upper Shelf 1

                                                Manufacturer – Sigma; Cat# T-6066

VERSENE

To make 5 L:

500 ml bottle of Dulbecco’s Phosphate-Buffered Saline

1 g EDTA for TC

4500 mls of bottled water

1. Mix and adjust pH to 7.1.

2. Aliquot into 100 ml bottle, put caps on LOOSELY and tape with autoclave tape.

3. Label with green tape (Versene, the date and your initials) and autoclave.

4. When bottles are completely cool, tighten caps.

Reagents Information:

Dulbecco’s PBS (10x): Location: Glasswares Cabinet

Manufacturer: Invitrogen/Gibco BRL; Cat# 14200-075

EDTA: Location – Lower Shelf 1

Manufacturer – JT Baker, Inc.; Cat# 4040-01

SECTION 14 Use of Influx and equipment in other laboratories

Influx Instructions for Operation –

Set-up of the Influx

1. Turn on the air conditioner (temperature thing by the main door to the lab, just push the On/Off switch, it should turn on). The cooling system and lasers make the temperature fluctuate a lot while in operation. This temperature fluctuation can have major effects on your drop delay. You need to keep the temperature of the room at a steady state.

2. Turn on the vacuum. It’s located in the wall above the lab bench. There’s a small lever that you need to turn to open (open position is at 6:00, closed position is at 3:00 and 9:00).

3. Turn on the air compressor for the table (Jun-Air box that the printer is sitting on top of) – generates a lot of heat, so that’s another reason why we had to turn on the air conditioner.

4. Sonicate the nozzle.

If the nozzle has been removed from the nozzle assembly, it can be found in an eppendorf tube in a 50mL tube in a blue tube holder– there should be 2 – 70µm nozzles and 2 – 100µm nozzles. The holder is on the shelf above the phone.

If the nozzle has not been removed from the nozzle assembly, carefully remove it by grasping the top of the assembly (the brown plastic part) and gently unscrewing the black hatched nozzle holder on the bottom of the assembly. It may help to brace your top hand holding the assembly against the black metal surrounding the nozzle assembly.

You will have just removed the small black piece from the nozzle assembly. This contains three parts –

· The black hatched outer part

· A small o-ring inside

· The actual nozzle (white part)

Using tweezers, take only the nozzle and place it into the sonicator (located on the low desk). There should be a solution of 10% NOX and 1% bleach in ddH20 in the sonicator, if not, you can fill it. (The bottle is located on the shelf above the low desk) If the solution looks grubby or thick, dump it out, rinse the sonicator and refill with fresh solution.

Turn the green switch on the front all the way to the right and the sonicator will start up. If it doesn’t, press the reset button on the top of the sonicator and then turn it to High again.

5. Start some water boiling on the hot plate in the lab (generally 100mL is enough).

6. While the nozzle is sonicating, you can go about filling up the sheath tank (Dandy).

Normally, we fill the tank with two days worth of fluid and empty it when we empty out the waste tank every second evening. You want to make sure the sheath fluid is at room temperature when you are running the machine so it’s best to keep the sheath fluid out on the shelf rather than in the fridge.

Generally the sheath and air lines will be connected to the rinse tank (Beau) or in a tube taped to the table leg, they need to be switched to the sheath tank. To disconnect the lines from Beau: from the ‘red’ line (sheath) hold onto the hatched bolt and lift up to release the lock and then lift the line from the joint. To connect this line to Dandy, once again lift up the hatched bolt on the line as you slide in the line. Once it is all the way in release the hatched bolt. To disconnect the blue line (air) hold down the hatched bolt and lift the line up from the joint. To reconnect this line hold down the hatched bolt on the base of the tank and slide the line in, once you hear a click, release the hatched bolt.

Also, whenever pressurizing Dandy or Beau, make sure the purge valve (the bolt between the air line and the pressure gauge) is closed by turning it in a clockwise direction.

7. Make sure all the valves are closed on the sorter head. There are three – sample (closest to the front), sheath (towards the back, the high valve), and vacuum (towards the back, the lower valve). These valves are found on the right side of the sorter head. They are most likely all left in the open position. Tighten them all.

8. Turn on the vacuum and air pressure. Flip the two switches on the left of the pressure controller up (the closest to the front is the pressure, the second one is the vacuum).

The final pressure of Dandy should be 28psi, so once you’ve flipped the switch make sure it is pressurizing. If it isn’t, check the line connections, the pressure release valve, and if necessary the seal.

Once it is pressurized (at least above 20psi), place the drain basin (the black basin connected to a pink line) under the nozzle and open the sheath line. This is to get any particles and a lot of the air bubbles out of the line without having them go through a cleaned nozzle. Let the fluid run until most of the air bubbles are out and then close the sheath line.

9. Open Spigot on the computer on the left. It will ask you if you’ve turned the box on already. At this point turn on the electronics box below the table. The button is on the bottom right of the box. You need to make sure when you do turn it on that the oscilloscope shows 3 lines. If it doesn’t, turn it off, wait a few minutes and turn it on again until you get the 3 lines. Once you have the oscilloscope showing three lines, you can then click OK on the Spigot electronics prompt.

10. Remove the nozzle from the sonicator when it is finished to flush it out (it will turn off automatically).

Using a syringe without the needle (found in the beaker next to the sonicator), suck up some boiling ddH₂O (don’t forget about the water you stuck on the hot plate to boil – this is what you want to use). Place the syringe end onto the pointed end of the nozzle (so you are aiming backwards into the nozzle) and blast out about half of your water. (**You always want to blast out the big end of the nozzle first just in case there is any debris still sitting in the nozzle casing.) Then, reverse the placement and shoot water the right way through the nozzle. If the spray is straight, the nozzle is clean. If it’s crooked and goes off in any sort of weird direction, have a look at it in the microscope, there’s gunk in there. Go back to step 4 and re-sonicate the nozzle tip.

Carefully place the nozzle tip back in to the black hatched holder, you may want to use the tweezers in the beaker by the sonicator. Make sure the o-ring is pressed all the way to the bottom (use the wooden end of a cotton swab to push it down if it’s getting caught up). The nozzle should be pushed all the way in also so it should sit straight in the holder.

Screw it back onto the nozzle assembly, being careful to hold the assembly at the last moment so it doesn’t twist.

11. Place the small drain basin (black U-shaped thing connected to tubing and just sitting on the table) under the nozzle assembly – this will catch any stray spray as you’re setting things up. If it’s not connected to anything, remove the rubber tubing and tip from the metal piece on the pink line and use that tubing for the drain basin.

12. Open the sheath and sample valves to fill them with fluid. Then turn off the valves, sample first. There will be a stream for a little while after you close the valve due to air bubbles in the nozzle. We need to get those out of there.

13. Fill the funny shaped tube with wide basin on the end of it (found in the beaker next to the sonicator also (looks like a mini-urinal)) with boiling ddH₂O and place it under the nozzle and immerse the nozzle end into the liquid.

14. Open the vacuum valve and watch the air bubbles get sucked out of the nozzle and into the vacuum tube. Keep an eye on the water level in your basin, you don’t want to run out or you’ll have to start all over again with filling the nozzle with fluid. While the vacuum is on and most of the bubbles are out, turn on the sheath line to loosen up any bubbles that may be there. Turn the sheath line and then the vacuum off. Open the sheath line again to once again loosen any bubbles that may be hiding in the Y junction. Close it and then turn on the vacuum line to check that there are no visible bubbles coming through the line.

To check for bubbles, close the vacuum valve and remove your basin from the end of the nozzle. Clean the end of the nozzle with a cotton swab (just the drops that sort of linger up there). Open the sheath valve and close it again. Does the stream stop pretty instantaneously, ie. does it stop before you have completely tightened shut the sheath valve?? If it doesn’t, there are still some air bubbles in the nozzle. Repeat step 14. (This is where having the electronics on helps, since you can look at the camera on the left to see when the stream shuts off) Once bubbles have been cleared, remove the drain basin.

15. Open the sheath valve fully and turn on the piezo to get rid of the spraying- on the sorter base flip up the switch labeled piezo next to the knob with red tape on it and then turn it clockwise to get a clean stream. On the camera screen on the right you should see the stream’s drop formation. If you do not see anything or the image of the stream is out of focus turn the little black vertical knob that is on the sorter head in front of the nozzle assembly and to the left. It is important to open the sheath line fully because if not then the drop formation will not be stable, open it to the point where the screw almost comes out.

Make sure the stream is hitting the drain hole in the center. If it’s not, use the positioners to adjust it. You might want to bring it into focus using the positioner at the back first and then adjust it side to side or angled.

A suggested method to focus the laser is:

i. Align the stream with the center of the length of the drain hole using the knob that faces out toward you.

ii. Align the stream with the center of the width of the drain hole using the knob on the upper right half of the nozzle assembly.

iii. Bring the nozzle tip into the video image on the screen on the left using the knob on the top of the assembly that points up.

iv. Bring the stream into focus using the knob behind the assembly pointing towards the window. The stream is in focus when the two sides of the stream are clean crisp well-defined lines.

v. Align the stream with the three pinholes using the knob on the lower right side of the assembly.

vi. With each adjustment, it’s possible that something will shift so repeat any of i. thru v. to get everything in alignment simultaneously.

16. Open the sample valve and let some sheath fluid backflush out – this serves two purposes, it cleans out anything that might be in there and it fills it up with fluid so there are no air bubbles that might make their way into the nozzle, and screw up drop your formation.

17. *** NEVER OPEN THE VACUUM VALVE!! ***
IT IS ONLY USED FOR CLEARING OUT AIR BUBBLES FROM THE NOZZLE!!

18. Slide the yellow cover in front of the nozzle area and replace the black top cover over the nozzle area.

19. Turn on the lasers – you can probably turn this on while you’re waiting for all the air bubbles to clear out of the nozzle. Check what the customer is staining with to see which lasers you will need.

a. UV laser –

i. If you need UV, turn on the cooling water.

ii. Open the chilled water supply valves – Make sure you open the valves in this order: RETURN (left), then SUPPLY (right). (There will be a lot of back pressure built up into the system if you turn on the supply first or forget to open the return.)

iii. Turn on the water cooler (right below the cooling supply valves). (power button on the front left side) You will hear it turn on.

iv. Flip the switch on the laser power supply underneath the table. Press the power button on the bottom left corner of the remote control. There is a delay when starting up the laser and the remote will show the countdown.

v. Open the aperture on the laser head (top hole on the actual laser)

vi. Crank the laser up to the proper wattage (the orange buttons on the right) – I usually run it at 150mW. That’s the middle reading on the remote control.

vii. Inevitably, you will have to tune the UV laser (when you can’t get any mW even when you crank it up to 45, but it shouldn’t be too far off. Squeeze your way to the backside of the table, remembering to put the remote somewhere that you can reach it from back there. Increase the amps to 45.

1. Hit “Power track” to turn it OFF,

2. Hit “Tune” to turn it ON.

3. There will be a bar that shows the maximum value that you hit while tuning it. Turn the adjusters on the back of the laser head one at a time until you’re maxed out on wattage.

4. When you’ve maxed out, hit “Tune” to turn it OFF.

5. Hit “Power track” to turn it ON. Make a note of your maximum wattage and then turn down the amps until you get about 150mW.

b. 488 laser –

i. power supply below the table has an on/off switch and a key. Turn both to the On (-) position – the cooling fan will turn on when you turn on the power.

ii. The Discharge button on the remote control can be turned to ON. And the standby button can be turned to RUN.

20. If you know the name of the protocol you are going to use, you can “Open a config file” from the File drop down menu, protocols are stored on the desktop in a folder called ‘Protocols’. If you don’t know, there is a protocol called UVSetup.sco that you can pull up to calibrate and make sure everything is aligned on the machine. You must open up the first protocol of the day twice to ensure that it opens properly with the proper settings.

21. Make sure you are triggering off the right signal. You need to look at the electronics box underneath the table. The Pre-amps are labeled and the ADC’s are labeled. The trigger goes into the “Signal Trigger” splitter and then comes back out and goes into the proper ADC.

i. UV-excitation: usually involves DAPI or Hoechst; use DAPI or HO into the 450-40 (linear signal) for trigger.

ii. 488-excitation: usually involves looking to see if something is dye + or dye -, usually with logarithmic signals; use Forward scatter for trigger.

iii. There are a few exceptions, check inside front cover of Protocols folder.

To check that you have everything connected properly you can check a file called ‘Protocol List.xls’, this file is stored in the “User Log” folder which has a shortcut to it on the desktop of the Summit computer. Find the protocol you are using on the ‘Search’ worksheet and you can check filters, pre-amps, adc’s, gains, and PMT settings.

Table 1: Normal ADC and Pre-amp assignments as of 9/25/03

ADC Assignments	Pre-amp Assignments

ADC 1 = 488RA scatter	Pre-amp 1 =
ADC 2 = 580/30 (PE)	Pre-amp 2 = 525/30
ADC 3 = Fwd scatter	Pre-amp 3 = 450/40
ADC 4 = 525/30 (FITC)	Pre-amp 4 = 615/40
ADC 5 = DNA integral (450/40)	Pre-amp 5 =
ADC 6 = DNA peak (450/40)	Pre-amp 6 = 580/30
ADC 7 = 615/40 (PI or EB) integral	Pre-amp 7 =Fwd scatter
ADC 8 = 615/40 (PI or EB) peak	Pre-amp 8 = 488 RA scatter
ADC 9 =	Pre-amp 9 =
ADC 10 = UV RA scatter	Pre-amp 10 =
	Pre-amp 11 = UV RA scatter
	Pre-amp 12 =

22. Before running someone’s samples run beads thru the machine to make sure that everything is in proper working order and everything is aligned. You may want to make adjustments to the focus of the nozzle assembly. To sharpen the focus of the lasers on the stream adjust the side positioner knobs on the laser heads on top of the sorter head. When focusing, generally first focus the laser that you will be triggering off and then any secondary lasers. To see if it is focused look at the Spigot screen and at a window displaying respective events- you want the population to be as tight as possible and as high as possible.

Note: Have your calibration beads running (when you can see them in the pinhole camera) before you switch to the ADCs/Trigger window in Spigot to change the trigger level and timing. See the Tricks to Spigot section at the end of these instructions for further info.

When taking a list always collect the following events: pulse width, ADC11, and ADC 12. Collect any other events that will be necessary for analysis. For your calibration lists, take a list of 10,000 events. Customer events will generally be larger (commonly: 20K, 30K, 50K, or 100K)

When saving files that you use for setup the following notation is used:

modyobjCustomerInitials

So for beads being run to setup an experiment for John Doe on Dec. 8^th the filename is: 1208bdsJD

Files are saved in: U:\FlowData\Influx and in the corresponding year, and month folders.

23. Make sure your event timing is correct if you are using multiple lasers.

On the oscilloscope, there is a 3^rd line at the top that shows the event window timing. There are three windows that represent the 3 pinholes and ultimately the 3 lasers, if necessary

In the ADCs/Trigger window, make Ch1 a 1^st event and Ch2 a 2^nd event by selecting an appropriate ADC Number. Then, use the event delay to adjust the timing of the 2^nd event with respect to the 1^st event.

In this window, you will also want to change the event order on the ADC boards.

i. For example, if you are setting up for a standard dual UV and 488 laser experiment,

1. ADCs 1&2 = event 2

2. ADCs 3&4 = event 2

3. ADCs 5&6 = event 1

4. ADCs 7&8 = event 2

5. ADCs 9&10 = event 1

*** Tricks to Spigot ***

When you are opening the first protocol of the day:

Start your calibration beads running first. Once you can see the beads fluorescing in the pinhole camera screen on the left, then check to see that Spigot is set up correctly. Go to the “ADCs/Trigger” window. It is likely that the trigger level will be higher than you want and in the Channel windows random ADCs will be chosen. Adjust the trigger level accordingly, generally for UV excited we run at 80-100, for Forward Scatter trigger level is between 200-300 and as high as 500; if you are unsure where to place the trigger, check the “Protocol List” file found in U:\FlowData\Influx\User Log\. Then select two ADCs to look at in the oscilloscope, if using two lasers look at ADCs for two different laser events. Adjust your timing if necessary. Go back to the sort window. If necessary adjust the PMT levels while tuning. Either write the original values down before changing, or you can reference the “Protocol List” to get these values right before you run their samples. These PMT settings are only starting points for running someone’s samples. You may have to change it, but generally the settings will be pretty close to perform an accurate analysis.

Some analysis protocols have sort windows drawn to give percentage values to the customer. Many times when opening a protocol with sort windows, all dots will be yellow, they should be either white, green, red, or some combination there of. If you have sort windows drawn, just modify their size slightly. You must do this to a left and right sort window. If you only have one sort window, draw the opposite window and then reset it to clear it in addition to resizing the sort window of interest and everything should be okay then.

If you are going to be sorting, hit the Bypass radio button and then the Normal radio button again for your sort windows. Spigot currently seems to be internally set to Bypass, and so if beads are falling in the sort windows then it will start sorting them and may make a mess down below.

Running your samples in the cytometer

For beginners:

Close the sample valve and put a tube into the holder. Be careful not to pull off the rubber plug or to crack the bottom of your tube. You can angle the tube into the holder and then place it vertical. Turn the red knob so that the pointer is facing towards you, NOT UP, then open the sample valve.

**Make sure when turning the red knob that you pull it out (away from the cytometer) as you turn it. **

Order of knobs when putting sample in and out of the system:

1. Close the sample nut to stop the back flushing on the sample line

2. Put the sample tube into the holder

3. Turn the red knob to point towards the front (it should’ve been pointing upwards)

4. Open the sample nut

5. Press the “BOOST” button on the pressure gauge assembly to blast your sample through. Watch on the pinhole camera and you can see when your sample is passing through the laser path

6. Take a list and make a printout if desired

7. Close the sample valve

8. Turn the red knob to point up

9. Remove the sample tube carefully – make sure the plug doesn’t go with it; if it starts to come out with the plug, push the tube all the way up and turn the red knob to face you. Then carefully pull the tube down, watching the rubber plug, it might still come out. The red valve provides a vacuum that will sort of pull it back on to the cytometer

10. Open the sample valve for back flushing, allow sample to back flush for approx. 30 seconds.

For more advanced users (recommended):

To load a sample, carefully and quickly with the tube slightly angled push it all the way up and then straighten it and bring it back down into the holder. It should have sealed in that process.

To remove a sample leave the red knob pointed towards you. Leave the sample nut open. Carefully and quickly, pull out your sample tube. You will need to push up and then down. As soon as you break the vacuum seal, the sample tubing will begin to backflush.

Let the system backflush for about 30 seconds before placing the next sample in.

** The reason for doing it this way is because the red knob is faulty and if used too many times can result in failure to create a vacuum seal on the sample itself. **

Shutting down the Influx

Lasers –

UV Laser

· Ramp down the UV laser on the remote.

· Turn OFF on the remote. (Off button on bottom left of remote)

· Underneath the table, turn OFF the big Coherent Innova Power supply.

· On the laser, close the UV aperture.

488 Laser

· Flip 488 laser to Standby (middle switch) on the remote.

· Flip OFF switch (switch on the left) on the remote.

· Underneath the table, flip OFF the switch and turn the key on the little box above the UV power.

Red Diode Laser

Unplug the laser from the power source behind the sorter head.

Fluidics –

· Fill the rinse tank (Beau) with about 200ml of DDH₂0. Occasionally you will want to boil it. Some times you will want to run ethanol instead to make sure everything is really clean. 200ml lasts about 20 minutes.

· Close the Sample setscrew.

· Close the Sheath setscrew.

· Flip Air pressure valve (closest to you on the pressure console) down.

· On the sheath tank there is a pressure release valve. You can open this slowly to quickly release the pressure in the tank.

· Once the pressure is completely released, use the quick releases on the two tubing pieces that go into the tank to release them from the sheath tank (Dandy).

· Insert them into the corresponding valves on the rinse tank (Beau).

· Re-pressurize (switch closest to you) and check the gauge on the tank to see that it goes to 28.

· If it doesn’t, flip switch and re-check tube-tank connections and the pressure release valve.

· Open the Sheath setscrew full.

· Open the Sample setscrew full

· Load a tube of ~10% bleach & water and run for 5-10 minutes and then remove the tube making sure that you allow the sample line to back-flush.

· Place the Erlenmeyer flask under waste drain.

* It is recommended to place the small black drain basin under the nozzle. The vacuum makes a distinctive sound when there is fluid being drained; you can use that sound to gauge when there is no more fluid in the sheath line because you will no longer hear the sucking sound.

Electronics –

· Turn off Piezo.

· In Spigot, go to the PMTs screen and turn off all the PMTs (hit the POWER buttons).

· Turn off the electronics box under the table or leave it on until the end to know when there is no more fluid running through the lines.

· Close Spigot.

· Turn off both monitors.

Lasers (revisited)–

The UV will have been running with the laser power off for about 10 minutes. Try not to let it run too long with out the laser power on, it builds up condensation.

· Turn off water cooler (LaserPur 40 cooler)

· Turn off water supply (closest to the door)

· Turn off water return (closest to you)

Fluidics (revisited)–

· Remove bleach/water tube after 5-10 minutes and let backwash.

· Wait for all the water to run through the sheath line. You’ll know when it does this by looking at the camera monitor or listening for the vacuum pulling through the small black drain basin.

· Once water stops, open the Vacuum setscrew valve. (All the valves will be open now)

· Turn off the electronics box below the table (switch on the lower right) if you haven’t already done so.

· Let the vacuum run ~5 minutes or until sheath (clear tubing) line is dry (frosted over).

· Flip both switches down on the pressure console.

· Turn off the air compressor (green button).

Miscellaneous –

· Close vacuum valve (over lab bench with red tape)

· Turn off AC.

· If you turned on the sample heater/cooler, make sure it is off.

· If you used the microscope, turn it off.

· If you used the sample-heating block by the printer, turn it off.

· Make sure the side door is closed.

· Turn off lights.

· Make sure the monitors of the two computers by the back wall are off.

PROTOCOL TO CALIBRATE MULTICHANNEL PIPETTES (Finnpipettes)

Software is in U:\PUBLIC\pipette calibration.

The most important thing is to make sure that the handle matches the block. When you open the latch of the handle the screen flashes with ‘calibration’ and it makes you select the type of block that you are going to put in. Make sure that the number of channels in the block and the maximum volume are correct. Then proceed to calibrate according to the software instructions:

1. Install the calibration software typing

SETUP on diskette drive.

2. Start the calibration program double clicking the icon.

3. First choose the volume range. Then enter the serial number

printed on the back of the handle e.g. K12345.

4. Enter current calibration factors to OLD FACTORS -fields.

To display current calibration factors do following steps:

First push down MODE-button and keep it down,

then push + and - -buttons as well. When all three buttons

are pushed down Release MODE-button first, then - and + -buttons.

Now CALIBRATE-text is blinking, push SET-button to accept.

Now current HK-factor is blinking followed by PK-factor.

Enter the factors to OLD FACTORS -fields

5. First do 10 pipettings at both calibration volumes,

then calculate the results and enter the mean volumes.

Then press ENTER or click CALCULATE-button.

6. Display the calibration factors again and change them with

+ or - buttons. Accept the new factor with SET-button.

7. Check the accuracy again and recalibrate if necessary.

The values for Finnpipette 12 channels 50ul are 14 163

The values for Finnpipette 8 channels 10ul are 10 132

The values for Finnpipette 8 channels 300ul are 13 141

Operation of GAMMACELL 40 Irradiator

H - 142 HSB

DEPARTMENT OF BIOLOGICAL STRUCTURE

Protocol from M. Lee REVISED 7 - 26 - 94

Only those individuals who have taken an instruction session from Andy Farr 206-685-1584 will be allowed to use the Gammacell 40.

1. Appointments must be scheduled in advance with Biological Structure, Cancer Research Lab., I - 534 in person. Schedule the Gammacell 40 in 1-hour blocks of time. Please do not reserve the Gammacell 40 between 12 noon and 1:30 pm. This time will be kept open for those who need to use it on short notice. Arrangements can be made for night and weekend use.

2. At the appointed time, the room key and irradiator key may be checked out from the Cancer Research Lab., I-534. Check the reservation book to be sure there are no scheduling conflicts. Each user will be required to sign for the keys in the book provided. Please list date, name of user, department, telephone number, grant name and number to which charges will be made, time keys taken, and time keys returned. Only authorized users whose name and signature are on file will be given the keys.
Please do not send someone to pick up the keys for you.

3. The Gammacell 40 is a self-shielded ¹³⁷ Cesium irradiator and no special precautions are necessary before entering H-142. A mechanical/electrical interlock system will not
allow the sources to leave the "safe" position when the sample cavity door is open, nor allow the door to open when sources are in the "on" position.

4. Turn on the Gammacell 40. An air compressor will turn on and run until the air storage
tank is filled. The irradiator will operate once sufficient air pressure is present in the system. This usually takes only a few seconds.

5. Set the appropriate switches on the control panel for operation:

6. "Manual/Auto" to "Auto" if using the timer.

7. Set timer by holding in the timer reset pushbutton located on the

8. left side of the windows while setting the timer with the push-

9. button switches below each digit window. Once the timer is set,

10. pressing the timer reset button will reset the timer for repeat

11. irradiations.

12. Turn on "Chamber Air" if needed. This supplies fresh air to the sample cavity for animals.

13. Open the sample cavity by pressing the sample drawer lock pushbutton and
pulling the door open.

14. Place experimental material in the sample drawer. Samples should be placed in the center
of the chamber where the irradiation field is most uniform. Close the sample door gently. For irradiating mice, use a holder designed to use for this machine.

15. Press "Source On" switch to start irradiation.

16. The sources will return to the "safe" position automatically when the timer has run out.

17. If the "Manual/Auto" switch is set on manual, press the "Source Off" switch to stop irradiation.
If it is necessary to stop irradiation at any time, press the "Source Off" switch. If irradiation
is stopped in the middle of an automatic timing cycle, pressing the "Source On" switch will
restart the timer with the remaining time.

18. When the irradiation is complete, open sample cavity door and remove experimental material.

19. To repeat irradiation procedure, simply press the timer reset button, reload the sample
cavity, and press "Source On" switch.

20. It is recommended that you leave the room during irradiations.
Remember to take the room key with you! The radiation level near the sample cavity
door is higher than background (about 5 mrem/hr). It makes sense to avoid any unnecessary exposure.(You would have to stand in front of the sample cavity door for 40 hours a week
for 25 weeks to reach the yearly occupational limit).

21. When finished, please clean up after yourself.

22. Wipe down sample cavity and holders with disinfectant provided. Leave sample cavity
door in closed position to decrease stress on the hinge due to the weight of the door.
Reset timer to zero. Turn irradiator off and remove key.

23. Return keys to I - 534. Record time keys returned in book.

All frequent users of the Gammacell 40 must wear a film badge. A film badge can be
obtained from the Radiation Safety Officer, GS-05, 543-0463.

It is recommended, but not required, that users of the Gammacell 40 take the radiation
safety course offered by Radiation Safety.

A copy of the users manual for the Gammacell 40 is located near the machine in H-142.
All users should be familiar with this document and should refer to it should questions arise.

Deviation from the prescribed rules and procedures, especially as it may affect safety or
the experimental work of other users, may lead to the termination of the privilege to
use the Gammacell 40.

EMERGENCY PHONE NUMBERS

NAME DAY NIGHT/WEEKEND

Andy Farr 685-1584 525-0928

Rick Hudson 543-0463

(Radiation Safety Officer)

If you are unable to reach one of the above at night or on weekends,

a Radiation Safety Officer may be contacted by calling University Police.

GAMMACELL 40 DOSE RATE SCHEDULE

Initial Calibration: 136.67 rad/minute on 12/15/85

¹³⁷Cesium half life 30.2 years

Dose rate decreases at 0.19 rad/min/month

Dose rates are posted at H-142, HSB (1 Gy = 100 rads)

Amsco Autoclaves

The autoclave is a large pressure cooker not unlike the one your mother (or you for that matter) might use for canning jams, jellies, etc. As such the same precautions should be taken with it as you would with a pressure cooker.

Basic Common Sense Rules.

1. Do not interrupt the cycle part way through unless the autoclave is alarming -- then press reset-- DO NOT OPEN THE DOOR- There is extreme danger of being scalded if you do this.

2. Do not try to force the autoclave open-the door should open easily if it doesn’t there is a problem

3. Do NOT attempt to sterilize flammable liquids

-there is a severe explosion risk

4. The most common injury with the autoclave is a burn - always wear protective gloves!

-even before starting a cycle the autoclave may be hot from a previous user

BE CAREFUL!

5. Always vent the autoclave for the full ten minutes at the end of a liquid cycle

-pressure differences between the inside and outside of the bottle can cause an explosion

1. Autoclave biohazard waste in the "dirty" autoclave only

Autoclave courtesy

1. Replace the paper roll when it is empty- making sure you put it in the correct way, otherwise nothing will print on it.

2. Sign in

3. If you remove someone else’s items, do it carefully.

4. Do not let your items sit in the autoclave for more than an hour after the cycle is finished.

5. If you change the setting on the autoclave return them to the setting listed on the wall after your cycle is finished

Using the autoclave

` There are two clean autoclaves and one dirty autoclave on the -1 level of K-wing. Pay attention to which autoclave you use, there are differences between them! All biohazard and only biohazard should be autoclaved in the dirty autoclave located in K-092 (across from the Fausto lab). Everything else should be autoclaved in one of the two autoclaves in K-059 (by the bathrooms)

Sterilizing pipette tips, microfuge tubes, paper or glass -

¨ In either of the two clean autoclaves place boxed pipette tips, or microfuge tubes on rack of autoclave or in an autoclave safe tray (i.e. nalgene, or metal) in the autoclave

¨ For paper, wrap in aluminum foil before autoclaving

¨ Any openings in the glassware (such the top) should be covered with aluminum foil to preserve sterility after autoclaving. Glass bottles with lids should be autoclaved with the lid loose to prevent explosion, make sure the tops are autoclaveable before attempting sterilization.

¨ Any autoclaveable plastics such as nalgene should also be autoclaved on cycle one. Be sure that all caps are loose to avoid distorting the shape or causing explosion.

¨ All items should be marked with autoclave tape before the cycle is started.

¨ Close the autoclave tightly, then activate cycle 1 by pressing it twice.

¨ At the end of the cycle inspect autoclave tape, if it has not turned black seek assistance.

Sterilizing solutions

Do not attempt to sterilize flammable solution! Filter sterilize them instead

¨ Place the bottle or flask containing the solution in a autoclaveable tray with a little (about an inch) of water in the base of the tray. Vent any caps, or cover flask opening with aluminum foil.

¨ The liquid cycle that should be activated is dependent on the volume that is being sterilized and the autoclave that is being used. In the gravity autoclave (the autoclave on the left in K-059) cycles three and four are for liquids. Cycle three is for volumes less than 250mls per bottle. Cycle four is for volumes between 250 and 1000mls. The vacumatic has only one liquid cycle, cycle four. It can sterilize volumes up to 1.5 liters. Any autoclave setting that will sterilize a larger volume than you are autoclaving will sterilize a smaller volume also, however those cycles take longer to finish.

Biohazard

¨ All autoclaving of biohazard materials in the K-wing should be done in the autoclave in K-092. No other autoclave should be used to sterilized biohazard wastes. Never run the biohazard autoclave overnight. The autoclave in K-092 has three cycles however for our purposes only the first two cycles will be used. Cycle one should be used to autoclave biohazard bags that will withstand only 121ºC. All other biohazard should be autoclaved using cycle two (132ºC). Bags with biohazard waste should be sealed with a rubber band to ensure proper venting. To autoclave biohazard waste put bags in plastic trays in K-092 with a little water in the bottom of the tray. A steri-strip should also be placed in the autoclave to guarantee the autoclave is sterilizing properly. Steri-strips are located in a small bag taped to the left side of the autoclave. The steam level must also be set to agree with the cycle. Cycle one requires a low steam setting, cycle two a high one. To change the steam setting open the door located at the top of the autoclave. There is a valve wheel that has high/low written on it. Turn the wheel in the direction indicated to change steam setting. Close the autoclave tightly and activate appropriate cycle by pushing the corresponding button twice.

Troubleshooting

Before opening the door of an autoclave verify the temperature is 80 degrees or less AND pressure is ZERO

¨ To turn the autoclave alarm off press the reset button

¨ Why is the autoclave making a horrible hissing noise?
More than likely the problem is because the door is not closed tightly. Try tighten the door. If the door is indeed closed it is possible that the door gasket is no longer good, if this is the problem someone has to be called to replace it.

¨ Why is the autoclave alarming?
If using a cycle that requires steam (i.e. biohazard, liquids) was water placed in the tray? Was the steam setting on the correct high/low setting. If either are the case turn the autoclave off and then on to restart the system then rerun. For almost any other problem physical plant should be called.

¨ Why can’t I close the door?
In order to close the autoclave the door has to be completely shut on both sides otherwise the latches will catch and you will not be able to lock the door. Usually the problem is the left edge where the hinges are. If you shift the door so as to push the left side first the problem should be eliminated.

Why can’t I open the door?
First make sure the cycle has really finished. The messages can be confusing. When the cycle is complete the read-out should say one of two messages:
“complete” dry cycle
“open door 10 min before unloading” liquid cycle

Dealing with the finicky pH meter

At the beginning of each day or if you suspect something is wrong during the day, try the following things to make sure the machine is calibrated.

1. Unplug the pH meter and then plug it in again. This will recalibrate the machine.

2. Remove the electrode connector and screw on the BNC shortening cap that is attached to the back of the meter.

3. Press pH/mV button on the front of the pH meter

A. If the reading is 0±0.5 mV then the pH meter has calibrated correctly.

B. If the reading is anything else, unplug and re-plug the machine and test again.

4. Electrode test

4. Now “slope” the calibration

Rinse probe and place in pH 7 buffer.

Press Setup until Clear buffers appears, then press Enter.

Check the pH of fresh 7, 10, and 12 buffers.

If the standards are not correct then press Standardize. Look for Good Electrode display.

Repeat step C with pH 10 and 12.

Press pH/mV to return to measuring mode at any point in this process.

NOTE: Keep your hands away from the electrode cable while measureing

Fluorometer

Fausto Lab K-088

For help see Katy or Martin (Rabinovitch Lab) or Jean Campbell in the Fausto lab.

1. Sign up for time to use the fluorometer on the sheet by the fluorometer

2. Turn on the machine at least 15 minutes before use (2 switches)

3. Open the program by clicking the fluorocount icon, and insert disk

4. File- open

5. Choose file

6. Click on Plate icon – “read settings” - check that everything is set correctly

7. Click on Plate out icon - take off lid and place plate in tray snugly with A1 in the upper left hand corner of the tray

8. Click on Plate in icon

9. Click on Plate icon - “read plate”

10. Click on Read icon

11. After the plate has been read, click cancel

12. File - export

13. Choose to export to: Excel

14. Choose “plate”

15. Choose “template”

16. Under export directory, choose drive a: 3 ½ floppy

17. Data to export- choose current file (ex: 7, 1/1)

18. Next

19. Export

20. Plate out- remove plate

21. Plate in

22. Close program and remove disk

23. Turn off both switches and clean the area

Opening and Saving File in Excel

When opening the Excel file, choose “delimited” and “tab” and “other”

Be sure to save the file and a Microsoft Excel Workbook and NOT as a text file

Storm 840 Phosphorimager

This imager is located in K-080 and is a shared instrument for the Departments of Pathology and Microbiology.

Before use:

1. Make a reservation for your time on the imager. The sign-up sheet is in K-080. The key for the room in on our master key ring labeled imager.

2. Expose your radioactive or fluorescent gel to a Molecular Dynamics screen for a few hours to overnight. These screens are VERY expensive, always handle them with care and only touch the edges.

Use of imager:

1. Bring your screen and a Zip Disk to room K-080.

2. Fill out the log book the day of use.

3. The computer and imager should be ON all the time, but you may need to hit the return key to inactivate the screen saver.

4. Place the Molecular Dynamics screen into the scanner and close the lid.

5. Open the STORM 840 Scanner control by double clicking on the STORM icon.

6. Choose the proper scanner type (radioactive or fluorescent).

7. Highlight the areas for scanning. For a big sequencing gel highlight the whole area.

8. Then click the “scan” box on the upper right corner.

9. Enter a name for your image file, then click “save”.

10. The machine will start to work. It takes about 11 minutes to scan a big gel.

11. When the imager has finished, transfer your file to your Zip Disk. This is important because the data in the STORM will be deleted at the end of each month. Please delete your own files when you no longer need them.

12. Close the STORM control -- DO NOT “park scan head”.

13. Erase your Molecular Dynamics screen in the light box.

SECTION 15 Protocols No Longer in Use

Assay for Detection of Phosphorylated Histone H3 by Flow Cytometry

Protocol from R.J. Widrow in C. Laird Lab.

1. Harvest cells. Centrifuge for 7 minutes at 1000 rpm (all subsequent centrifugation steps are similarly performed).

2. Aspirate medium. Resuspend in 5 mls of PBS. Pellet cells.

3. Aspirate medium. Resuspend in 1 ml of PBS. Forcefully pipette cells into 5 mls of ice cold 80% ethanol. Incubate cells for 2 hours at -20°C.

4. Pellet cells. Resuspend in 5 mls of 1% BSA in PBS. Pellet cells again.

5. After centrifugation, resuspend cells in 1 ml of 1% BSA, 0.25% triton X-100 in PBS and place on ice for 5 minutes.

6. Add 5 mls of 1% BSA in PBS. Pellet cells.

7. Resuspend cells in 200 ml of PBS containing 1% BSA, 10% NGS. Add 0.5 mg of anti-phosphorylated histone H3 antibody (Upstate Biotechnology) per sample and incubate cells at 4°C overnight.

8. Add 5 mls 1% BSA in PBS. Pellet cells.

9. Resuspend cells in 200 ml of PBS containing 1% BSA, 10% NGS. Add 5 mg of FITC conjugated goat anti-rabbit IgG (Fab’2 fragment) per sample and incubate 1 hour at 4°C.

10. Add 5 mls of 1% BSA in PBS. Pellet cells. Resuspend in 500 ml DAPI.

Western Protocol for WRN Protein

From Monnat lab Kate, Yannick, and Christina (with Swisshelm Lab modifications)

Cell Lysis:

Trypsinize cells (if necessary), count and centrifuge desired number of cells 5-7 min at 200-300 x g, 4⁰C.
*Note: It is extremely important to keep everything cold from this point on (paranoia is fine).
Remove medium, and resuspend cells in 1ml cold PBS (non-sterile is OK).
Transfer cells to 1.5 ml microfuge tube (cold) and spin down for 5-7 min at 2000 rpm (300 x g), 4⁰C. Remove supernatant.
Cells can be snap-frozen at this point and stored at ^–80⁰C; thaw on ice and flick tube with finger to resuspend cells before adding lysis buffer. Do NOT vortex.

Prepare Lysis Buffer (50 ml):

(this high salt buffer is good for extracting proteins bound to DNA)

(final concentration)

ddH₂O 36.75 ml

1 M TRIS pH 8.0 1.25 ml (25 mM)

0.5 M EDTA 500 ml (5 mM)

5 M Na Cl 6.0 ml (600 mM)

glycerol 5.0 ml (10%)

100 mM DTT 5 ml (10 uM)

10% NP-40 (Nonidet P40) 500 ml (0.1%)

= Igepal

Add the following protease inhibitors (per ml of buffer) no more than 10 minutes before use (standard “cocktail” is also fine):

10mg/ml leupeptin 1 ml (10mg/ml)

10mg/ml pepstatin A 0.5 ml (5 mg/ml)

100 mM PMSF 10 ml (1 mM)

Add cold lysis buffer to thawed or fresh cells that have been resuspended by gentle flicking (see above). Pipet gently to completely resuspend pellet. It is important to use a volume of lysis buffer that is approximately 6x the volume of the cell pellet (20-30 ul for10⁶ cells seems to work well; 10⁶ cells gives good signal on Western). Note: after final centrifugation, DNA in pellet makes it difficult to recover the full volume of lysate, so start with more cells than you need.
Incubate lysate on ice for 40 min., flicking tube occasionally.
Centrifuge 30 min. at maximum speed (14K rpm / 16K x g for Eppendorf microfuge).
Label tubes for desired size aliquots and chill. (20ul aliquot is enough for quantitation of total protein). Transfer supernatants to tubes and snap freeze in liquid nitrogen or ethanol/dry ice bath. Store at –80^oC

Western (minigel):

Pour 7.5% gel (WRN protein runs at approximately 160kD) and 4.5% stacking gel. See BioRad/Swisshelm Lab protocols for running and transfer to PVDF membrane.
Thaw samples on ice; add appropriate volume 4x Sample Buffer; boil 5-7min and centrifuge briefly (if samples have been frozen after boiling, simply re-boil for 3 min. and centrifuge).
Load samples and MW markers and run dye front just off gel.
Transfer to membrane and continue with blotting protocol.

Immunoblotting (as in Swisshelm Lab protocols, but for this particular anti-WRN (Transduction Lab) primary Ab, an anti-mouse IgG₁-HRP must be used (Southern Biotechnolgy):

Incubate membrane 10 min in 10% H₂O₂ (optional step, but important if samples contain endogenous peroxidase).
Block 1-2 hours at RT in Blocking Buffer (2% milk powder in PBS/0.1% Tween 20) Note: Monnat Lab uses 10% milk and TBS/T and blocks O/N at 4^oC).
Incubate 1 hr at RT in 1^o Ab (Transduction Labs mouse anti-WRN monoclonal) at 1:1000 in Blocking Buffer
Wash 3x 5 min. in PBS/T
Incubate 45 min – 1 hr at RT in 2^o Ab (Southern Biotechnology Associates goat anti-mouse IgG₁ – HRP at 1:2000 in Blocking Buffer.
Wash 4x 5 min. in PBS/T.
Place wet blot on plastic wrap and cover with ECL reagent (Pierce Super Signal West Pico); Incubate for 5 min.
Drain blot slightly, wrap in plastic, place in cassette and expose to film.

Electroporation Guide

The specifics of each electroporation vary from experiment to experiment.

This protocol is an outline of general guidelines for electroporations.

D.D.Hickstein, E. Grunvald, G. Shumaker, D.M. Baker, A.L. Back, L.J. Embree. E. Yee, and K. A. Gollahon. Transfected leukocyte integrin CD11b/CD18 (Mac-1) mediates phorbol ester-activated, homotypic cell:cell adherence in K562 cell line Blood 82:2537-2545. 1993.

Before you begin (start to handle cells) do the following:

1. Outline the experiment defining cell lines and plasmids.

a. You will need about 10⁷ cells in culture for each cuvette/parameter

b. Make use you have an adequate amount of DNA -- 100 mg for each parameter

2. If you have enough cells and plasmid, then you can continue.

3. Set-up (takes 30-90 minutes)

a. Place one bottle of DMEM + 10% FBS + P/S + L-glut. at 37^oC.

b. Fill two buckets with ice.

c. Make sure there are pipettes 10’s and 25’s and tips IN the hood.

d. Label small petri plates to match the parameters in the outline for the experiment.

e. Make sure the HEPES buffered saline is ice cold.

f. Calculate the volume of each plasmid prep which you will need. This will vary so make sure this parameter has been determined in the outline. Make a list and tape it to the front of the laminar flow hood for easy reference. Copy this list into your notebook and create a column to record the time constant (T) for each cuvette.

g. Set 2 channels of a timer for 10 minutes, but do not start yet.

Actual electorporation

1. Make sure steps 1,2, and 3 above have been completed. These experiments are timed and there is little or no time to do these things once you have started the experiment,

2. General “RULES”.

a. Do not electroporate more than 10-12 cuvettes at time. If you need to do more than this number, break the experiment down into sets and complete one set before beginning the next one.

b. Once you begin handling cells, do not stop or delay any step.

c. Do not add DMEM to small petri plates until 1-2 minutes before plating cells. The media become too alkaline and this very hard on cells which have just been electrocuted.

3. Harvest the cell type to be used into a single flask and determine the concentration of cells by counting them on a hemacytometer.

4. Determine the volume of cells that you will need for this experiment (10⁷cells/cuvette) and harvest the appropriate volume in 50 ml tubes.

ex. Cell count is 1.2 x 10⁶ cells/ ml

You need 10⁷ cells / cuvette 10⁷^{=
8.3 mls / cuvette}

1.2 x 10⁶

If you will be using 5 cuvettes then you will need to harvest

5 x 8.3 mls = 41.5 mls. for this experiment.

5. Spin the cells in the Sorvall tabletop centrifuge at 1000 rpm (a setting of 3) for 8 minutes.

6. WHILE THE CELLS ARE SPINNING, label the sterile cuvettes to match the parameters being tested.

7. Remove the centrifuged cells from the centrifuge and pour off the supernatant into a bleach flask.

8. Resuspend the pellet in 35 mls of HEPES buffered saline (wash step). If you have several tubes, e.g., you spun down 100 mls, pool the cells into a single tube with 35 mls of HEPES buffered saline. Spin the cells at 1000 rpm for 8 minutes as above.

9. WHILE THE CELLS ARE SPINNING, rinse the cuvettes with HEPES buffered saline. Remove the saline with a vacuum line.

10. Remove the centrifuged cells from the centrifuge and pour off the supernatant into a bleach flask.

11. Resuspend the pellet in 0.5 mls of HEPES buffered saline for every cuvette to be used (final concentration 2 x10⁷cells / ml).

ex. If you will be using 5 cuvettes, resuspend the pellet in 2.5 mls.

12. Use the P-1000 and pipette 500 ml into each cuvette.

13. Now add the appropriate amount of DNA to each cuvette and cap it. Transfer the cuvette to an ice bucket immediately after adding the DNA. Start the timer (10 minutes) immediately upon placing the first cuvette in the ice bucket.

14. When the set of cuvettes is finished move the ice bucket next to the electorporator and set the machine for the appropriate mF and Volts for that cell line. The settings are on the top of the machine for each cell line.

15. When the 10 minutes are up, electroporate the cuvettes in the same order that they were placed in the bucket. Start the timer (10 minutes) immediately upon completion of electroporating the first cuvette. Place the cuvette in the second ice bucket and record the time constant for each sample.

16. When the timer is at 1-2 minutes, add 5 mls of warm DMEM+ FBS to each petri dish.

17. When the timer sounds, begin to plate the cuvettes into the petri dishes. Move complete set into C0₂incubators soon as possible after plating.

HEPES BUFFERED SALINE

1.2 g HEPES

                8 g NaCl

0.37 g KCl

0.1 g Na₂HPO₄

1.08 g glucose

pH to 7.4; qs to 1 liter with dH₂0; filter sterilize

Retrovirus Production

A.D. Miller and G.J. Rosman. Improved retroviral vectors for gene transfer and expression. Biotechniques 7:980-990. 1989

A.D. Miller, D.G. Miller, V.J. Garcia, and C.M. Lynch. Use of retroviral vectors for gene transfer and expression. Meth. Enzymol.217:581-599. 1993.

Viral Production for Transient Transfection

Day 1

1.Plate out PE501 cells at 5 x 10⁶ per 60 mm tissue culture dish.

Day 2

1. Using 15 ml polypropylene tube (orange cap) prepare Stock solution:

500 mM HEPES, pH 7.1 100 ml 50 mM Final

2 M NaCl 125 ml 250 mM

150 mM NaPO₄, pH 7.1 10 ml 1.5 mM

H₂O 765 ml

1 ml

2. Use 5 ml polystyrene (clear) tubes, prepare DNA solution:

DNA Vector 5 ml (concentration ~2 mg / ml want 10 mg, adjust H₂O)

2M CaCl₂ 25 ml

H₂0 170 ml

200 ml
3. Add 200 ml each stock solution to separately marked 5 ml polystyrene tubes.

4. Add 200 ml DNA solution dropwise to stock solution in 5 ml tubes (use P-200, takes around 2 minutes). Knock tube (like finger vortexing) while adding. Should see some cloudiness, if not something is wrong – DO NOT CONTINUE!

4. Let sit 30 minutes in hood.

5. Change media of the tissue culture dishes with 4 mls fresh DMEM + 10% FBS.

6. After 30 minutes of DNA precipitation, add whole 400 ml mix using P-1000. Drop around whole plate. Rock to mix. Put in incubator.

Day 3

1. Change media on tissue culture dishes.

Day 4

1. Harvest media from tissue culture dishes and centrifuge at 3,000 x g for 5 min to remove debris.

2. This media can be used to infect cells or be stored at -70 until needed

Solutions (Filter sterilize all solutions):

2 M CaCl₂ (FW= 147 for CaCl₂ 2 H₂0)

2.94 g qs to 10 ml with dH₂0

500 mM HEPES

Use 1 M HEPES stock for TC.

5 mls HEPES and 5 mls d H₂0 pH should be about 7.1

2 M NaCl (FW=58.44)

1.17g qs to 10 mls with dH₂0

150 mM Phosphate buffer (pH 7.0)

Monobasic NaH₂PO₄ H₂0 (FW= 137.99)

0.207 g qs to 10 mls with dH₂0

Dibasic Na₂HPO₄ (FW=141.96)

0.213 g qs to 10 mls with dH₂0

Mix about 4.5 mls of monobasic solution with 10 mls of dibasic solution. Check pH.

If pH is too high continue to add monobasic solution.

Infection of PA317 Cells

Day 1.

1. Transfect PE501.

Day 2

1. Plate out PA317 at 5 x 10⁵ cells/plate

2. Plates are 60 mm Falcon 3002 tissue culture plates.(DO NOT USE PETRI DISHES)

3. Change media PE501 transfections.

Day 3

1. Collect supernatant from PE501 transfections. Can store at -70^oC. Spin at 3000 rpm, 5 min.

2. Change media PA317 plates (mark also).

3. Add 10 ml, 100 ml, 1 ml PE501 sups to PA317 plates, respectively
(in duplicate).

4. Add 4 ml polybrene (stock 4 mg/ml, final 4 mg/ml)

Day 4.

1. Split transduced PA317 cells 1/10. (100 ml / 1 ml trypsin). Add to 4 mls DMEM

2. Add G418 to 1 mg / ml final (stock 100 mg /ml) to split cells

G-418 solution

1 Weigh dry powder. Should be 1g (for 1g size)

2. Add 9.7 mls DMEM (w/o FCS).

3. Filter through 0.2 micron (0.22 um) syringe filter.
[Geneticin G-418 Sulfate Invitrogen/Gibco # 860-1811, around 66% active]

Store in the refrigerator at 4^oC

Or Use pre-mixed Geneticin from Invitrogen/Gibco # 10131-019 50 mg/ml

Collection of Clones Infected with Retrovirus

1. Circle clones with marker.

2. Add 1 ml trypsin, rock 5 sec, and remove trypsin.

3. Add cloning rings (Bellco) to each clone using forceps. Firmly but gently make sure pushed down.

4. Add trypsin, around 75 ml with a P-200 pipette.

5. Wait until cells detach and ball up.

6. Add 1 ml DMEM with appropriate selection agent to 24 well plates. Mark plate.

7. With P-200. pipette, resuspend contents gently within ring. Add to individual wells.

When confluent

1. Add 500 ml PBS or trypsin to wash. Remove.

2. Add 500 ml trypsin. Wait until detaches.

3. Resuspend with p. pipette. Add whole 500 ml to 4 ml DMEM in 60 mm plate (can use less if other clones less confluent).

Viral Titers

Collect viral supernatant by:

Day 1. Split clone 5x10⁵ C in 4 mls.

Day 2. Change media.

Day 3. Harvest supernatant.

For TK infection.

Day 1. Split TK confluent cells to 5 x 10⁵ Cells.

Day 2. Remove sup. Add new 4 mls DMEM.

Add 1 ml (10 ml and 100 ml) viral supernatant

Add 4 ml polybrene.

Swirl to mix.

Day 3. Split 1/10 into a 60 mm TC dish and put the other 9/10 into a 100 mm TC dish. Add media with appropriate selection (ex G418)

Approximately 7-10 days later, Count clones:

1. Wash with 2 mls PBS.

2. Add 1 ml Coomassie blue stain.

3. Wait 1-2 minutes.

4. Rinse off in sink.

5. Clones are visible to the naked eye. Count and record.

6. Calculation of titer.

# of colonies x inverse of the dilution = titer in cfu/ml

ml of virus added

ex. For 77 colonies from 100 ml and a 1/10 dilution.

77 x 10 = 7700 cfu/ml

0.1

Coomassie blue stain: 1 g/liter of Coomassie Blue powder

40% methanol

10% acetic acid

Infection of Cells with Retrovirus

A.D. Miller and G.J. Rosman. Improved retroviral vectors for gene transfer and expression. Biotechniques 7:980-990. 1989

A.D. Miller, D.G. Miller, V.J. Garcia, and C.M. Lynch. Use of retroviral vectors for gene transfer and expression. Meth. Enzymol.217:581-599. 1993.

Day 1

1. Plate out cells to be infected at 5 x 10⁵ cells/plate

2. Plates are 60 mm Falcon 3002 tissue culture plates.
(DO NOT USE PETRI DISHES)

Day 2

2. Change media on all plates (4 mls/ plate)

3. Add 10 ml, 100 ml, or 1 ml of viral supernatant to individual plates

4. Add enough fresh media to each plate to bring the volume to 4 mls

5. Add 4 ml polybrene (stock 4 mg/ml, final 4 mg/ml)

6. Incubate overnight.

Day 3.

1. Trypsinize cells and plate 1/10 of the cells in one 60 mm TC dish, and 9/10 of the cells in a 100 mm TC dish..

2. Add to 4 mls DMEM with serum

3. Add G418 (or appropriate selection reagent) to 1 mg / ml final (stock 100 mg / ml) to split cells.

4. It will take 7-10 days before cells that did not take up the virus die.

G-418 solution

1 Weigh dry powder. Should be 1g (for 1g size)

2. Add 9.7 mls DMEM (w/o FCS).

3. Filter through 0.2 micron (0.22 um) syringe filter.
[Geneticin G-418 Sulfate Invitrogen/Gibco # 860-1811, around 66% active]

Store in refrigerator at 4^oC

Or Use pre-mixed Geneticin from Invitrogen/Gibco # 10131-019 50 mg/ml

Transformation of Lymphocytes

1. Adjust the total number of cells in each tube is know, pipette 2.4 - 4.8 x 10⁶ cells into each of four 15-ml tubes. This concentration will permit the distribution of 1 - 2 x 10⁵cells to each of 24 wells in a 96-well plate from each 15-ml tube.

2. Spin 10 min at 1000 rpm, aspirate off RPMI.

3. Add 2.4 ml Epstein-Barr virus(from fresh or refrigerated stock is best, but frozen stock can be used –just thaw on ice or at room temperature and then keep in refrigerator at 4^°C) to each tube. Use virus from 2 different dates (date 1 & date 2).

4. Incubate for approximately 2 hrs at 37^oC, or overnight up to 24 hrs.

5. Then take one tube (date 1) and add 2.4 ml (1:1 dilution) of 20 mg/ml phytohemagglutinin A (PHA, Sigma, St. Louis, MO) in RPMI + 16% FBS. The final concentration of PHA is 10 mg/ml. Mix very gently and pipette 0.2 ml into each of 24 wells on a flat-bottom 96-well plate. Fill the outer row of wells with sterile H₂0 to prevent evaporation.

6. To a second tube (date 1) add 2.4 ml (1:1 dilution) of 2 mg/ml cyclosporin A (CsA, Sandoz, East Hanover, NJ) in RPMI + 16% FBS. The final concentration of CsA is 1 mg/ml. Gently mix and pipette 0.2 ml into each of 24 wells as above.

7. Repeat with tubes from viral date 2.

8. Incubate at 37^oC, 5% CO₂, humidified incubator.

9. One to 2 days later examine cultures for evidence of transformation (transformed cells will appear blast-like and form clumps). You’ll notice that the cells treated with PHA will begin to grow rapidly. This can save weeks of time unless there is a T-cell-mediated immune response. In that case, the PHA-treated cells will die in massive numbers. With the CsA-treated cells the emergence of transformed clones is slower but failure after blastogenesis appears to be lower. Thus, these cultures are a valuable backup.

10. At about 2 weeks, feed the PHA wells with plain RPMI + 16% FBS. Feed the CsA wells with RPMI + 16% FBS + 1 mg/ml CsA. If there are nice healthy clumps of cells and no cell death, then supplementation with CsA can be discontinued, otherwise continue to add for at least 3 weeks.

11. Most cell lines can be subcultivated after a month. Many of these have good cell growth and can go directly into a 25-cm² flask. Other cell lines require careful screening of the wells. The best wells (least cell death, good growth) can be subcultivated into a 24-well plate, 2-5 little wells/big well. There they can hopefully outgrow the immune response, and be fed and split 1:2 wells as they grow, eventually being moved to a 25-cm² flask for mass culture.

12. After the culture is established, three separate freezings (cryopreservation) are carried out at 1-2 week intervals, at least 3 ampules each time. Two vials from separate freezings are stored in a back-up freezer system.

Freezing medium is cold RPMI + 20% FBS + 10% DMSO. There should be at

least 4 x 10⁶ cells/ml of freezing medium.

Notes

1. We use no antibiotics in the medium. In our experience, the presence of penicillin and streptomycin retards the frequency of successful transformation.

2. As CsA is an extremely hydrophobic substance, its solubility constitutes as important technical point for in vitro experimentation. The compound prepared as described below remains soluble in the concentrations mentioned in this paper. A stock of 1 mg/ml was prepared by dissolving 1 mg CsA in 0.1 ml ethanol , then adding 0.02 ml Tween 80 and mixing well. Finally RPMI 1640 medium was added, while continuously stirring, dropwise to a final volume of 1.0 ml. Further dilutions were made in this medium. Appropriate solvent controls were always included.

Arbitrarily Primed PCR (AP-PCR) old version

Ap-PCR detailed protocol revised 9/16/2004 by Nancy Linford

Arbitrarily Primed Polymerase Chain Reaction (AP-PCR) requires the use and understanding of several technologies prior to beginning. These include isolation of specific tissues by various techniques, DNA purification, preparation of ³²P labeled primers using T4 polynucleotide kinase, PCR, denaturing polyacrylamide gel electrophoresis, and data collection and analysis using the Molecular Dynamics Storm PhosphoImager system and ImageQuant Software.

Tissue Isolation

Before beginning AP-PCR, an understanding that DNA from different tissues is being compared for genomic instability, typically suspect tissue versus more stable constitutional tissue. In the case of ulcerative colitis (UC) or pancreatic cancer, epithelial tissue which is suspect for high genomic instability will be compared to constitutional tissue (e.g. stromal tissue). Therefore, the technique of tissue isolation is very important. Mixing of tissues can lead to mixed results since genomic instability is estimated at 5 to 15% in epithelial cells.

The technique of choice for separating epithelial cells from stromal cells is the EDTA Shake-Off Method. See the EDTA Shake-Off Method for details. Other sources of tissues include tissue culture, Laser Capture micro-dissection and other methods of micro-dissection. Once the tissues are isolated, then the DNA can be purified for analysis.

DNA Purification

DNA purification can be accomplished by several methods, such as the classical phenol/chloroform method, or via the use of spin columns. The method of DNA purification by using spin columns is best accomplished by following the manufacturers methods (e.g. see Qiagen). In house, the phenol/chloroform technique works very well. See the Phenol/Chloroform Method for details.

Once the DNA is purified, DNA concentration must be determined for each sample. This is accomplished by one of two standardized techniques. One, using a spectrophotometer, establish the absorbency at 260 nm and 280 nm for each sample. The standard conversion is one A₂₆₀ OD unit is equal to 50 ng/ml of DNA. To determine the purity of each sample, the A₂₆₀/A₂₈₀ ratio should be greater than 1.5. Another technique is to determine DNA concentration by using the Hoeffer Scientific Instruments Mini-Fluorometer (TKO 100). Follow the manufacturer’s instructions to determine DNA concentration for each sample.

With DNA concentration for each sample completed, the DNA’s are ready of PCR.
Primers: #1 AP-PCR-F 5’-AAC CCT GAC TGC CTT TCC-3’
#2 MCG1 5’-AAC CCT CAC CCT AAC CCC AA-3’
#3 BLUE 5’ CCG AAT TCG CAA AGC TCT GA-3’

Day 1 ³²P labeling of primers

Use filtered pipet tips and place used tips in acrylic container.
In a .5 ml tube combine (behind shield):

T4 polynucleotide Kinase	1 ml	Rack 7 box 1
10x PNK buffer	1 ml	Rack 7 box 1
³²P	2 ml	Hood (can use for 1 month)
Primer (APF,Blue,or MCG1) (100 ng/ml)	6 ml	Rack 7 box 2 – aliquot to avoid repeated freeze-thaw cycles

Place in Eppendorf thermocycler under “kinase” program

37°C 1 h

65°C 15 min

4°C hold

Add 15 ul of sterile water and 5 ul of cold primer
Store at -20°C in an acrylic shielded box
Monitor the area for radioactive contamination
Primers used: AP-F, MCG-1, and BLUE

PCR reactions

Always include a control reaction with No DNA to confirm that reagents are not contaminated.
Make the reagents up in order as a cocktail and add to DNA in 0.2 ml tube. Keep reagents on ice and ensure that they have thawed completely - vortex to mix
Do duplicate reactions of each DNA and always include the suspect DNA and the constitutional control in the same reaction set.
Once the primer is added, all tips must go in acrylic box and reactions must be behind shield.

Sterile Water	3.69 ml	Hospital grade
10xPCR buffer with MgCl₂	0.7 ml	Roche - Rack 7 box 1
MgCl² (25 mM)	0.4 ml	Roche - Rack 7 box 1
DNTPs (10 mM) *	0.15 ml	From Biochem stores – Rack 7 box 1
³²P Primer mixture	1 ml	-20°C (from above)
Taq polymerase	0.06 ml	Roche - Rack 7 box 1
DNA (10 ng/ml)*	1 ml	Rack 5 box 3

*aliquot to reduce freeze-thaw cycles

put sterile mineral oil on top of each reaction
use “AP-HOT” program on Mjresearch thermocycler or “APPCR” on eppendorf

Program:

94°C 3 min

Low stringency portion (5 cycles)

94°C 30 sec

50°C 1 min

72°C 1.5 min

High stringency portion (25 cycles)

94°C 15 sec

60°C 15 sec

72°C 1 min

72°C 7 min

4°C hold

Prepare Gel

Set prepared gel plate setup on the bench. Set the top end on saran wrap for neatness. Place the clear plastic adapter with grey foam over the bottom of the gel and secure it using the white knobs. You should be able to see the line between the plates clearly. Add the clear plastic adapter and hose found in the drawer beneath the eppendorf cycler.
In the hood (caution – neurotoxic acrylamide):

100 ml 6% gel solution

500 ml 10% APS (frozen in rack 8 box 1)

50 ml TEMED

Mix and pour into a beaker. Once these are added you have 15 minutes before the gel becomes too hard to use. Work quickly!
Suck up gel mix in 100 ml syringe. Get at least 80 ml.
Put end of syringe in tube (no air bubbles) and press out liquid with a single consistent motion.
Carefully remove tube and cap off end with parafilm
Use 60 ml syringe to suck up excess gel.
Place comb in the gel
Use remaining gel to create a bead across the top to prevent a reduction in well size when the gel dries and shrinks.

· Cover top with saran wrap and allow to polymerize overnight. Leave remaining gel in beaker to test polymerization

Day 2 Prepare the gel for loading

Carefully remove the comb while the gel is lying down. Peel off excess acrylamide.
Place in lower buffer chamber and attach electrodes. Fill lower buffer chamber with 1xTBE and upper buffer chamber with 1xTBE.
Wash out and straighten the wells

Prepare samples for loading

Mix each sample with 7 ml formamide loading dye (stored at RT behind gel apparatus)
Heat each sample to 95°C for 8 minutes then cool on ice
Wash the wells of the gel immediately before loading sample
Load 1.2 ml per lane
Run at 2000 volts for 6 hours

Drying the gel

Start the gel dryer (dark room in TC room) heating to 80°C
Take whole gel apparatus to the sink
Dump out the upper buffer chamber by attaching hose to attachment on side
Cap off hose and place in lower buffer chamber (radioactive) without losing suction
Bring gel apparatus to the bench and place with plain glass plate (gel slick coated) facing up
Remove upper buffer chamber. If all goes well the gel should be stuck to the bottom chamber
Take out sheet of 3mm paper and stick to the gel.
Trim off excess gel and place in radioactive waste.
Remove the gel/paper combo and cover with saran wrap.
Place in gel dryer at 80°C for 45 minutes (with vacuum on) – saran wrap facing up
Place in cassette with phosphor screen facing the gel overnight

Day 3 Scan the gel

· Remove the screen and throw the gel into the radioactive trash

· Go to Storm Phosphorimager (in room with ice) and place screen on scanner

· Scan entire screen and save to a file on the desktop

· Bring down a disk and transfer data.

Reset the screen

· Place screen on bright light box by phosphorimager for 10 minutes

Data analysis

· Open file in ImageQuant Tools

· Reduce the size and save it to the public folder of the hard drive.

· Open the file in Image Quant 5.0

· Use the pline tool to select a lane

· Preferences ® object attributes ® line width=3-5

· Shift-arrow to select multiple lines

· Object alignment – top (sets all lines to start on same pixel)

· Object ® object manager ® rename to identify lanes

· Bottom – click “create graph” icon

· Use magnifying glass to see how many bands are expected (30ish expected)

· Analysis ® peak finder ® baseline= lowest point, kernel = 20ish

· Click on the baseline and press insert to enter a point

· Drag baseline to obvious peak starts and ends. Be consistent between lanes

· Go back with zoom (at bottom – select area) and clean up baseline.

· Peak definition icon looks like a peak with 3 sides of a square. Click this

· Click at start of peak and drag to end.

· To delete peaks, click on page icon (at bottom) for area definitions

· Analysis ® area report ® report

· Double click to export into excel

· Copy into template.

· Ratio of % of total area is the value we are interested in

· Flag as unstable if >2.0 or <0.5

· Sum instability over total # of peaks.

ImageQuant Analysis Considerations

The important feature to remember for analysis is that the constitutional baseline values must be duplicated very closely for each epithelial sample from the same patient. Always work in sets of patient lanes. Set up the lanes for each patient group starting with the constitutional tissue first, then the epithelial lanes for easier comparisons. Compare each chromatograph with the gel itself to determine if a visible band is present. Work with between 20 and 30 well-isolated and distinguishable bands per primer. The total of bands used for analysis is between 60 and 90 bands. After determining the baseline and conducting a Peak Analysis, a Peak Report is conducted and then exported into Excel for final analysis. Use the % Peak values for analysis only. Look for a greater than 2X and less than 0.5X differences between bands of constitutional vs. epithelial lanes. Also, look for new or missing bands. These differences are scored as genomic instability. Determine the percent genomic instability by dividing the number of different or variable bands by the total number of bands times 100.

Inter Simple Sequence Repeat PCR (ISSR)

ISSR is a method of DNA fingerprinting which can be used to measure genetic differences between DNA samples. Purified DNA samples are used in the protocol listed below, then run on a denaturing acrylamide gel and PCR products are quantitated with ImageQuant software. Protocols for these other techniques (DNA purification, acrylamide electrophoresis, and PCR product imaging and analysis) are listed in the Arbitrarily Primed PCR (AP-PCR) protocol.

³²P labeling of Primers

Prior to running a PCR reaction, the primers must be 5’-end labeled with ³²P. Two different primers are used for ISSR PCR. These include primer #1 “CA8RG” with sequence 5’-CACACACACACACACARG-3’ and primer #2 “CA8RY” with sequence 5’-CACACACACACACACARY-3’. “CA8RY” is a 1:1 mixture of two separate primers, “CA8RT” and “CA8RC” (The "Y" sequence as defined in the protocol is correct; that is, "Y" = 1:1 of C:T).

The ³²P polynucleotide kinase reaction is as follows:

T4 Polynucleotide kinase 1 ml

10X PNK Buffer 1 ml

³²P 2 ml

Primer [100 ng/ml] 6 ml

1. Incubate at 37°C for one hour.

2. Incubate at 65°C for 15 minutes.

3. Cool on ice, then add 5 ml cold-primer [100 ng/ml] and 10 ml dH₂O.

Total of 30 ml per reaction; use 1 ml for each 7 ml PCR reaction.

PCR Conditions

Once the DNA concentrations have been determined and primers ³²P-labeled, the PCR reactions are now ready to be conducted. Prior to setting up the reactions, setup a thermal cycler with the following program:

1. 94°C 3 minutes

2. 94°C 30 seconds

3. 55°C 45 seconds

4. 72°C 2 minutes

Repeat steps 2 – 4 29X for a total of 30 cycles

5. 72°C 7 minutes

6. 4°C hold

Setup each PCR reaction as follows:

10X PCR Buffer with 25 mM MgCl₂ 0.7 ml

20% Formamide 0.7 ml

0.1% Triton X 0.7 ml

dNTP’s (25 mM) 0.06 ml

P-32 Labeled Primer [30ng/ml] 1.0 ml

Taq Polymerase 0.06 ml

dH₂O 2.78 ml

DNA [10 ng/ml] 1.0 ml

7 ml

After each PCR reaction is prepared, place the samples into the thermal cycler and run the program. After the thermal cycler program has completed add 7 ml formamide loading dye to each sample of a hot reaction run. Samples should be stored at -20°C in an acrylic box suitable for storage of radioactive materials.

Acrylamide (6%) gel solutions for ISSR and AP-PCR

Caution: Liquid Acrylamide is Toxic. Blue Nitril gloves and a lab coat should be worn when handling.
All Acrylamide needs to be polymerized before disposal.

6% Acrylamide Gel Mix

75 ml Acrylamide 40% Stock (19:1) (Acrylamide: Bis-Acrylamide, 19:1)

50 ml 10X TBE

200-250 ml dH₂O

225 g Urea

Stir in a graduated cylinder until Urea goes into solution (>1 hour).

Adjust final volume to 500 ml with dH₂O.

Filter to remove any particulates, standard filter paper without any vacuum works well.

Store in a brown glass bottle (acrylamide is photosensitive) at 4°C.

Gel mix should be used within 1 month of preparation date.

Gel Polymerization

Have the sequencing plate prepared for pouring the gel before initiating polymerization.

For a 38cm x 50cm sequencing gel, use ~100 ml Acrylamide gel mix.

Add 500 ml 10% APS (Ammonium Persulfate Solution) to the gel mix.

Add 50 ml TEMED to the gel mix and stir briefly.

Pour the gel immediately after preparing for polymerization.

Allow at least 1.5 hours before using to allow complete polymerization.

Leave any unused acrylamide solution in the beaker or flask to confirm that the gel solution polymerizes

ABI sequencing (PCR product) Protocol.

1. Run PCR reaction, make sure to have at least 100ng of product, usually 50ml reaction should be enough.

2. Use QIAQUICK SPIN kit to purify PCR product, elute DNA into 20ml of ddH₂O.

3. Big dye reaction:

10ml purified PCR product (>= 50ng, see note1)

8 ml Bigdye mix

2 ml primer (= 4 pmol, use either forward or reverse primer)

20ml reaction

After mixing, promptly put into PCR machine for the following cycling:

95^oC 5 min

(95^oC - 3 sec; 50^oC - 10sec; 60^oC - 4min) 30 cycles

4^oC

4. Clean up the above reaction. If available, please use commercial clean up kit,

otherwise we could use isopropanol for clean up.

Isopropanol protocol:

1) Transfer PCR product to 1.7ml tube, then add 80ml of 75% isopropanol.

2) Vortex to mix, then room temperature for 15 min.

3) Spin high-speed 16k rpm for 20 min.

4) Discard supernatant.

5) Add 250ml of 75% isopropanol

6) Vortex briefly

7) Spin high speed (16k rpm) for 5 min.

8) Remove supernatant.

9) Vacuum dry.

5. Now ready to send over to sequence facility. The dry sample can be stored at –20^oC for up to two weeks.

Note 1: if the PCR product is bigger than 500 bp, need to increase PCR amount
2: after the big dye reaction, should avoid light exposure.

TdT with no Antibody Stain

Ref. Li, X. Traganos, F. Melamed, M.R., and Darzynkiewicz, Z. 1995. Single-step procedure for labeling DNA strand breaks with fluorescein- or BODIPY-conjugated deoxynucleotides: detection of apoptosis and bromodeoxyuridine incorporation. Cytometry 20:172-80

A. Cell Fixation

1. Collect cells from culture or Rx into 15 ml conical tubes, spin down, 1200 rpm (~200
    g) for 10 minutes. Resuspend in »1ml PBS. (If labeling with mAb, go to Section D
    and do not fix cells until after mAb labeling step).

2. Add volume of 2% paraformaldehyde equal to the amount of PBS in the tube.

3. Fix in the refrigerator (4°) for 15 minutes, avoid over fixation.

4. Add PBS to about 10 ml (rinse)

5. Centrifuge at 800-1200 rpm (~200g) for 15 minutes.

6. Carefully remove the supernatant and resuspend in 300 ml of PBS.

7. Add ice cold 95% EtOH dropwise (3 volumes of EtOH to 1 volume of cells) while vortexing. ex. If your cells are in 300 ml of PBS, then add 900 ml of EtOH. (final concentration: about 8x10⁵ cells/ml). Can be stored for at least 72 hrs. at -20°C.

B. Assay

1. Add »0.5-1x10⁶ fixed cells to siliconized, serum treated microfuge tubes. Add PBS to
1.5 ml. Centrifuge 15 min @ 1200 rpm.

NOTE: Always set up two thymus or HL-60 controls, ± »1mM camptothecin for ~ 3 hrs, one for the reaction + DAPI stain and one for the reaction only (e.g. FITC only). Also set up another DAPI only tube for the cells of interest to check two things, first is the "machine" working correctly and second to determine the level of FITC autofluorescence.

2. Remove supernatant (to about 100 ml). Add 1 ml of normal saline. Centrifuge 15 min
@ 1200 rpm.

3. Remove supernatant to exactly 100 ml. Split into two 50 ml aliquots: ½ gets 50 ml of
the TdT reaction mix with TdT enzyme and the other ½ gets 50 ml of the TdT reaction mix without TdT enzyme.

TdT reaction mix with enzyme for dUTP FITC				TdT reaction mix with enzyme for dUTP-BODIPY
Reagent	per tube	_____tubes	Reagent		per tube	_____tubes
5X TdT buffer*	10.000 ml	_______ml	5X TdT buffer		10.0000 ml	_______ml
CoCl₂(25mM)	5.000 ml	_______ml	CoCl₂(25mM)		5.0000 ml	_______ml
BSA (20mg/ml)*	5.000 ml	_______ml	BSA (20mg/ml)		5.0000 ml	_______ml
dUTP-FITC(0.78 mM)	0.125 ml	_______ml	dUTP-Bodipy(1mM)**		0.0625 ml	_______ml
TdT enzyme (25U/ml)	0.400 ml	_______ml	TdT enzyme (25U/ml)		0.4000 ml	_______ml
dH₂0	30.000 ml	_______ml	dH₂0		30.0000 ml	_______ml

Total Volume	50.000 ml	_______ml	Total Volume		50.0000 ml	_______ml

*5X TdT buffer: 1M sodium cacodylate, 125mM Tris-HCl pH 6.6 @ 4^o, 1.25 mg/ml BSA, 10mM CoCl₂ (comes with enzyme from Boehringer Mannheim. BSA is endonuclease free (Boehringer Mannheim).

** Molecular Probes, Inc. Cat#C7614

I. Vortex. Incubate 1.5 hours @ 37^oC

II. If you are doing ONLY the TdT assay:

A. wash one time with 1 ml of 15mM EDTA + 0.1% NONIDENT P-40 (IGEPAL CA-60) + 0.1mM 2-ME (300 ml of 500mM EDTA + 10 ml of NONIDENT P-40 (IGEPAL CA-60) per 9.7 ml of water)

B. If the cells were labeled with mAb and put through the TdT assay wash with 1 ml of PBS + 10mM EDTA. (mAb binding may be EDTA sensitive so verify 10mM is not too conc.)

III. Spin and remove supernatant.

IV. Add 200 ml DAPI (5 mg/ml). Run up to 24 hr later.

C. Treatment of microfuge tubes

1. Set up 50 to 100 1.5 ml microcentrifuge tubes in a rack.

2. Add 0.5 ml of Sigmacote (Sigma SL-2) to the first tube, remove this 0.5 ml from the first tube, and add it to the second tube, repeat with 3rd, 4th, 5th etc.

3. Air dry.

4. Add 0.5 ml of fetal bovine serum to the first tube, remove this 0.5 ml from the first tube and add it to the second tube, etc.

5. Rinse with 1 ml fresh PBS in each tube.

6. Air dry and place in sealed bag.

D. Variation for immunofluorescence.

Step 0. Label cells with mAb and wash with PBS.

Fix cells up through step A5. Replace PBS in step A6 with PBS + 0.1% NONIDENT P-40 (IGEPAL CA-60). Skip EtOH step #A7.

Can be stored 24 hr at 4°C. Wash one time in saline. Start TdT labeling at step B.3.

After TdT labeling is best to run same day; probably OK 24 hr later (stored 4°).

E. Variation for 7AAD Viability +/- Immunofluorescence

Before fixation, add 2.5ug/ml 7AAD to cells, incubate 30 min on ice, rinse with PBS

Proceed as in step A.2 with paraformaldehyde fixation, then NONIDENT P-40 (IGEPAL CA-60) permeabilization.

F. Control

A background fluorescence -TdT enzyme control is very desirable. Substitute the TdT enzyme mix with a solution of 1mM EDTA, 4mM 2-ME, 50% glycerol (v/v), 200 mM KCl, pH 6.5.

TdT Assay for HPBL

Rabinovitch Lab - 8/8/95 -

I. HPBL Antibody Labeling

A. Mark 100 ml level on siliconized, serum treated microfuge tubes.

B. Place control, -TdT control, and treated HPBL, ³ 10⁶ cells, in separate labeled microfuge tubes.

1. Eight (8) additional aliquots of ~0.5-1X10⁶ cells will be needed for FACS controls (AKA “machine” controls), see page 3.

2. Centrifuge the samples 15 minutes @ 1200 rpm.

3. Carefully remove the supernatant to 100 ml.

C. Label (stain) samples with mAb (also see F.2.a, below):

1. Add 28½ ml Ab mix to each sample.

2. Re-suspend cells with pipette.

D. Incubate on ice for 30 minutes.

E. After Ab incubation, add 1 ml PBS to each sample.

F. Re-suspend cells with pipette.

G. Centrifuge samples for 15 minutes @ 1200 rpm.

H. Carefully remove the supernatant to 100 ml.

per sample	X ____ samples = _____ tubes	Source
5 ml aCD4-PerCP	X _____ tubes = _______ ml aCD4-PerCP	___________________
4 ml aCD8-PE	X _____ tubes = _______ ml aCD8-PE	___________________

20 ml PBS	X _____ tubes = _______ ml PBS
Total:	X _____ tubes = _______ ml Ab mix

I. HPBL Fixation

A. Add 400 ml cold (4°) PBS to each sample (A.4, above).

B. Re-suspend cells with pipette.

C. Add 500 ml cold (4°C) 2% paraformaldehyde (PF) to each tube
(i.e., add a volume of 2% PF equal to the volume in tube {~500 ml} for 1% PF fixation).

D. Incubate at 4° for 15 minutes; avoid over-fixation.

E. After the PF fixation, dilute each sample with ~300 ml cold (4°) PBS (i.e., fill to ~1.4 ml).

1. Centrifuge 15 minutes @ 1200 rpm.

2. Carefully remove the supernatant to 100 ml.

C. HPBL permeabilization

C.1 Add 1 ml of PBS with 0.1% NONIDENT P-40 (IGEPAL CA-60) to each sample.

C.1.a Re-suspend cells with pipette.

C.1.b. Centrifuge 15 minutes @ 1200 rpm.

C.1.c. Carefully remove the supernatant to 100 ml.

C.2 Add 1 ml saline to each sample.

C.2.a Re-suspend cells with pipette.

C.2.b. Centrifuge 15 minutes @ 1200 rpm.

C.2.c. Carefully remove the supernatant to exactly 100 ml.

D. TdT Assay (plus see E., below)

D.1 Split samples into two 50 ml aliquots.

D.2 Label 1 50 ml aliquot with TdT reaction mix (not the -TdT controls, i.e., see E., below)

D.2.a Add 50 ml of the TdT reaction mix (see table below) to each TdT+ sample.

D.2.a Re-suspend cells with pipette.

D.2.b Incubate for 1½ hours @ 37°C.

D.3.Add 1 ml of 10 mM EDTA in PBS. (Verify that this EDTA conc. does not remove your mAb!)

D.3.a. Centrifuge 15 minutes @ 1200 rpm.

D.3.b Carefully remove the supernatant to 100 ml.

TdT reaction mix for dUTP-FITC			TdT reaction mix for dUTP-BODIPY
Reagent	per sample	____ samples	Reagent	per sample	____ samples
5X TdT buffer¹	10.0 ml	______ml	5X TdT buffer¹	10.0 ml	______ml
CoCl₂(25mM)¹	5.0 ml	______ml	CoCl₂(25mM)¹	5.0 ml	______ml
BSA (20mg/ml)²	5.0 ml	______ml	BSA (20mg/ml)²	5.0 ml	______ml
dUTP-FITC³	0.125 ml	______ml	dUTP- BODIPY⁴	0.065 ml	______ml
TdT enzyme (25U/ml)¹	0.40 ml	______ml	TdT enzyme (25U/ml)¹	0.40 ml	______ml
dH₂0	30 ml	______ml	dH₂0	30 ml	______ml

-TdT Control reaction mix for dUTP FITC			-TdT Control reaction mix for dUTP-BODIPY
Reagent	per sample	_____ samples	Reagent	per sample	_____ samples
5X TdT buffer	10.0 ml	_______ml	5X TdT buffer	10.0 ml	__________ml
CoCl₂	5.0 ml	_______ml	CoCl₂	5.0 ml	__________ml
BSA	5.0 ml	_______ml	BSA	5.0 ml	__________ml
dUTP-FITC	0.125 ml	_______ml	dUTP-BODIPY	0.065 ml	__________ml
mock enzyme¹	0.40 ml	_______ml	mock enzyme¹	0.40 ml	__________ml
dH₂0	30 ml	_______ml	dH₂0	30 ml	__________ml
Total Volume	50.0 ml	_______ml	Total Volume	50.0 ml	__________ml

2. Boehringer Mannheim (5X buffer: 1 M sodium cacodylate, 125 mM Tris-HCl {pH 6.6 @ 4°C}, 1.25 mg/ml BSA, 10 mM CoCl₂)

3. Boehringer Mannheim (endonuclease-free BSA)

4. Boehringer Mannheim (Fluorescein-12-dUTP, 0.78 mM).

5. Molecular Probes, Inc., Cat. No. C7614 (1 mM)

D.4. Add 200 ml DAPI (5 mg/ml) with 10% DMSO.

D.5 Re-suspend cells with pipette.

D.6 After TdT labeling, it is best to run same day; OK to run up to 24 hours later (store at 4°C).

E. No TdT Enzyme Control (AKA -TdT Control) and FACS Controls

E.1 TdT controls (to determine sample background fluorescence): Use mock enzyme in the reaction mix.

E.1.a Treat the -TdT controls from D., above, with 50 ml of the following rxn. mix:

E.1.b Then proceed as in D.4, above.

1. Solution of 1mM EDTA, 4mM 2-ME, 50% glycerol (v/v), and 200 mM KCl, pH 6.5.

Annexin V Assay for HPBL

Rabinovitch Lab - 8/8/95

A. HPBL DNA (Hoechst Dye 33342) Labeling

A.1 Mark 100 ml level on siliconized, serum treated microfuge tubes.

A.2 Place control and treated HPBL in separate labeled microfuge tubes

A.2.a Use ³ 1X10⁶ cells per sample

A.2.b Eight (8) additional aliquots of ~0.5-1 x 10⁶ cells will be needed for FACS controls
(AKA “machine” controls), see E, below.

A.3 Centrifuge the samples 15 minutes @ 1200 rpm.

A.4 Carefully remove the supernatant to 100 ml.

Hoechst mix per sample	X ____ samples = _____ tubes	Source
1 ml RPMI + 10% FBS	X _____ tubes = _______ ml RPMI
10 ml Hoechst 33342 (100X)	X _____ tubes = _______ ml Hoechst	_____________
Total:	X _____ tubes = _______ ml mix

A.5 Stain samples with the following mix (i.e., not the FACS controls; see page 3):

A.5.a Add 1.01 ml of the mix to each sample.

A.5.b Re-suspend cells with pipette.

A.5.c Incubate at 37°C for 30 minutes.

A.6 After Hoechst 33342 incubation:

A.6.a. Centrifuge samples for 15 minutes @ 1200 rpm.

A.6.b Carefully remove the supernatant to 100 ml.

B. HPBL Antibody Labeling

B.1 Label (stain) samples from A.6, above, with mAb (also see FACS controls, E, below)

B.2.a Add 28½ ml Ab mix to each sample.

B.2.b Re-suspend cells with pipette.

B.2.c Incubate on ice for 30 minutes.

per sample	X ____ samples = _____ tubes	Source
5 ml aCD4-PerCP	X _____ tubes = _______ ml aCD4-PerCP	______________
4 ml aCD8-PE	X _____ tubes = _______ ml aCD8-PE	______________
0.3 ml Ho 33342	X _____ tubes = _______ ml Hoechst 33342	______________
20 ml PBS	X _____ tubes = _______ ml PBS
Total:	X _____ tubes = _______ ml Ab mix

Annexin Binding Buffer without EDTA

10 mM HEPES

140 mM NaCl

2.5 mM CaCl₂

10 ml Hoechst 33342/ml buffer

B.3 After incubation, add 1 ml ice-cold 1X Annexin V Binding Buffer to each sample as follows:

B.4. Re-suspend cells with pipette.

B.5 Split each sample (leaving tubes on ice)
(i.e., put ½ of each sample into another siliconized, serum treated microfuge tube, marked “-”):

B.5.a ½ (550 ml) for Annexin without EDTA (remains in original tube) (positive sample); and

B.5.b ½ (550 ml) for Annexin with EDTA (negative sample).

B.5.c Centrifuge samples for 15 minutes @ 1200 rpm.

B.5.d Place samples on ice.

B.5.e Carefully remove the supernatant to 100 ml, put samples back on ice.

C. HPBL Annexin V Staining

Annexin mix per sample	X ____ samples = _____ tubes
1.25 ml 1X Annexin V-FITC	X ___ samples = ___ ml 1X Annexin V-FITC
3.75 ml Binding Buffer	X ___ samples = ___ ml Binding Buffer
Total:	X ___ samples = ____ ml mix

C.1 Add 5 ml ice-cold of the following mix to each 100 ml positive and negative sample (kept on ice):

C.2 Use 1X Annexin V-FITC (not the 10x stock often provided in kits.)

C.3 Re-suspend cells (kept on ice) with pipette.

C.4 Incubate on ice for exactly 10 minutes.

Annexin V with EDTA samples (AKA Annexin negative samples {A-})	Annexin V without EDTA samples (AKA Annexin positive samples {A+})
10 mM HEPES	10 mM HEPES
140 mM NaCl	140 mM NaCl
1 mM EDTA	2.5 mM CaCl₂
10 ml Hoechst 33342/ml buffer	10 ml Hoechst 33342/ml buffer

C.5 After incubation, add 1 ml ice-cold Annexin Binding Buffer as follows:

C.5.a Centrifuge samples for 15 minutes @ 1200 rpm.

C.5.b Place samples on ice.

C.5.c Carefully remove the supernatant to 100 ml.

D. HPBL Fixation

D.1 Add 300 ml 4° 1% paraformaldehyde with 1% Hoechst 33342 to each tube (C.3, above).

D.1.a i.e., add 1% PF with 10 ml Hoechst per ml

D.1.b Re-suspend cells with pipette.

D.2 Store at 4°, OK to run up to ~24 hours later.

E. Annexin FACS controls (AKA “machine” controls): HPBL Antibody Labeling

E.1 Mark 100 ml level on siliconized, serum treated microfuge tubes.

E.2 Place eight (8) aliquots of ~5X10⁵ - 10⁶ HPBL in separate labeled microfuge tubes.

E.2.a Four (4) FACS controls to be labeled here:

E.2.a.i PE only (two {2} samples: one will become PE only, one will become PE + FITC);

E.2.a.ii PerCP only; and

E.2.a.iii PE + PerCP.

E.2.b Four (4) FACS controls to be labeled later:

E.2.b.i No stain;

E.2.b.ii FITC only;

E.2.b.iii FITC + Hoechst 33342; and

E.2.b.iv Hoechst 33342 only.

E.3.Centrifuge all eight (8) FACS controls 15 minutes @ 1200 rpm.

E.4 Carefully remove the supernatant to 100 ml.

E.5 Label one PE + PerCP FACS control with the following mix:

E.5.a 4 ml aCD8-PE

E.5.b 5 ml aCD8-PerCP

E.5.c 20 ml PBS

E.6 Label two (2) PE only FACS controls with the following mix:

E.6.a 4 ml aCD8-PE

E.6.b 25 ml PBS

E.7 Label one PerCP only FACS control with the following mix:

E.7.a 4 ml aCD8-PerCP

E.7.b 25 ml PBS

E.8 Add 1 ml PBS to the remaining four (4) FACS controls.

E.9 Re-suspend cells with pipette.

E.10 Incubate on ice for 30 minutes.

E.11 After incubation, add 1 ml PBS to the PE + PerCP, PE only, and PerCP only FACS controls.

E.11.a. Centrifuge all eight (8) FACS controls for 15 minutes @ 1200 rpm.

E.11.b. Carefully remove the supernatant to 100 ml.

F. Annexin FACS controls: HPBL Fixation

F.1 Add 400 ml cold (4°C) PBS to all eight (8) FACS controls, re-suspend cells.

F.2 Add 500 ml cold (4°C) 2% paraformaldehyde (PF) to each FACS control.
(i.e., add a volume of 2% PF equal to the volume in tube {~500 ml} for 1% PF fixation).

F.3 Incubate at 4°C for 15 minutes; avoid over-fixation.

F.4 After the PF fixation, dilute each tube with ~300 ml cold (4°C) PBS (i.e., fill to ~1.4 ml).

F.4.a. Centrifuge 15 minutes @ 1200 rpm.

F.4.b. Carefully remove the supernatant to 100 ml.

F.5 Dilute the following four (4) FACS controls with 200 ml PBS and hold in refrigerator (4°C):

F.5.a PE only;

F.5.b PerCP only;

F.5.c PE + PerCP; and

F.5.d No stain;

F.6 Hold the DAPI only FACS control in refrigerator (4°C) (i.e., no dilution).

G. Annexin FACS controls: HPBL permeabilization

G.1 Add 500 ml of PBS with 0.1% NONIDENT P-40 (IGEPAL CA-60) to the following three (3) FACS controls:

G.1.b FITC only;

G.1.b PE + FITC; and

G.1.c FITC + DAPI.

G.2 Re-suspend cells.

G.3 Centrifuge 15 minutes @ 1200 rpm.

G.4 Carefully remove the supernatant to 100 ml.

G.5 Add 1 ml saline to the samples and the three (3) FACS controls, G.1, above, and re-suspend cells.

G.5.a. Centrifuge 15 minutes @ 1200 rpm.

G.5.b. Carefully remove the supernatant to 50 ml.

H. Annexin FACS controls: TdT Assay

H.1 Treat the three (3) FACS controls from G.5, above, with the following reaction mix:

-TdT Control reaction mix for dUTP FITC			-TdT Control reaction mix for dUTP-BODIPY
Reagent	per sample	___ samples	Reagent	per sample	_____ samples
5X TdT buffer	10.0 ml	________ml	5X TdT buffer	10.0 ml	__________ml
CoCl₂	1.5 ml	________ml	CoCl₂	1.5 ml	__________ml
BSA	5.0 ml	________ml	BSA	5.0 ml	__________ml
dUTP-FITC	0.125 ml	________ml	dUTP-BODIPY	0.065 ml	__________ml
mock enzyme¹	0.40 ml	________ml	mock enzyme¹	0.40 ml	__________ml
dH₂0	33 ml	________ml	dH₂0	33 ml	__________ml
Total Volume	50.0 ml	________ml	Total Volume	50.0 ml	__________ml

1. Solution of 1mM EDTA, 4mM 2-ME, 50% glycerol (v/v), and 200 mM KCl, pH 6.5.

H.2. Add 50 ml of the TdT reaction mix to each of the three (3) FACS controls.

H.2.a Re-suspend cells with pipette.

H.2.b Incubate for 1½ hours @ 37°C.

H.3. Add 1 ml of 1 mM EDTA in PBS to each of the three (3) FACS controls.

H.3.a. Centrifuge 15 minutes @ 1200 rpm.

H.3.b Carefully remove the supernatant to 100 ml.

H.4. Add 200 ml DAPI (5 mg/ml) with 10% DMSO to the following two (2) FACS controls.

H.4.a DAPI only (stored at 4°C); and

H.4.b FITC + DAPI.

H.5. Add 200 ml PBS to the following two (2) FACS controls.

H.5.a FITC only; and

H.5.b PE + FITC.

H.6 Re-suspend cells with pipette.

Acridine Orange - Alkaline Unwinding DNA Damage Assay

1. Before harvesting cells

1.1. Pre-cool on ice or hold at 4°C:

1.1.1. 0.15N (0.9%) (150mM) NaCl, pH 7.4 (verify pH)

1.1.2. 0.15N (0.9%) (150mM) with 0.1% Triton X-100, pH 7.4 (verify pH)

1.1.3. 150mM TRIS buffered 0.9% NaCl (TBS) pH 7.5 (verify pH)

1.1.4. 40% & 30% ethanol {probably optimal EtOH conc., higher conc. increases variation}

1.2. Coat ½ dram glass vials with 30 ml 0.1% LGT agarose & dry in 65^oC heat block (approx. 60’)

1.3. Melt 2.25% ULGT in 65^oC heat block; when melted hold @ RT-37^oC

2. Harvest cells at end of drug treatment or prior to H₂O₂ or gIR treatment

2.1. Optimal cell concentration approx. 0.25-1X10⁶/ml (1X10⁶/ml runs quicker on cytometer)

2.1.1. may have problem w/ cell-to-debris ratio in final cell suspension when <0.1X10⁵ cells/ml initially

2.1.2. >2X10⁶ cells/ml initially may exceed optimal cell-to-mM AO ratio in final cell suspension

2.2. Re-suspend the cells in serum-free DMEM with 1.0 mg/ml nuclease-free BSA

2.3. Hold on ice until <4°C (approx. 10min.) before fixation {>30’ on ice may increase variation}

3. H₂O₂ Treatment

3.1. Treat cells on ice at approx. 1X10⁶/ml DMEM/BSA

3.2. H₂0₂ Rx (10 or 100 mM H₂0₂ 5-10min. sufficient as assay control for most cells)

3.2.1. 0.03% (8800 mM) H₂0₂

3.2.1.1. 10 ml 30% commercial H₂0₂ in 10 ml distilled water

3.2.1.2. 1.14ml 0.03% H₂0₂ per ml cell suspension for 10mM final increment (e.g., 5.7ml for 50mM final)

3.2.2. 0.003% (8800 mM) H₂0₂

3.2.2.1. 1 ml 30% H₂0₂ in 10 ml distilled water

3.2.2.2. 11.4ml 0.003% H₂0₂ per ml cell suspension for 10mM final increment (e.g., 57ml for 50mM final)

4. H₂O₂ Treatment with Repair

4.1. Treat cells on ice per above

4.2. Centrifuge at 4°C @ 800-1000 rpm 3-8 min. (time in centrifuge counts as treatment time)

4.2.1. For 5 min. treatment, centrifuge approx. 3 min.

4.2.2. For 10 min. treatment, centrifuge approx. 8 min.

4.3. At end of treatment/centrifuging, drain and re-suspend at 1 x 10⁶/ml in DMEM/BSA

4.4. Fix No Repair Rx & No Repair controls immediately

4.5. Incubate remaining samples in 37° water bath (or incubator) for the desired length of repair time

5. 5. gIR Treatment

5.1. gIR Rx (1 to 10 Gy [1Gy = 10 rad] sufficient as assay control for most cells)

5.2. Treat cells on ice at approx. 1X10⁶/ml DMEM/BSA

6. 6. gIR Treatment with Repair

6.1. Treat cells at approx. 10X10⁶/ml DMEM/BSA

6.2. At end of treatment, add 100ml cell suspension to 900ml 37° media with BSA

6.3. Fix No Repair Rx & No Repair controls immediately

6.4. Incubate remaining samples in 37° water bath (or incubator) for the desired length of repair time

7. Fixation

7.1. Add 3 ml ice-cold 40% EtOH drop-wise while slowly vortexing {EtOH conc. will affect variation}

7.2. Using tubes coated with 1:20 BSA:PBS may reduce cell loss

7.3. Fix on ice for 10min. minimum

8. Wash (skip & go to 9., below, for endonuclease treatment)

8.1. Centrifuge approx. 800-1000 rpm 10-15min.; drain supernatant

8.2. Re-suspend pellet in 0.5 ml ice-cold 30% ethanol

8.3. While slowly vortexing, add 5 ml ice-cold 0.15N NaCl drop-wise {solubilizes excess protein}

8.4. Centrifuge 800-1000 rpm 10-15min.; drain supernatant

9. Endonuclease Treatment (if desired)

9.1. Following EtOH fixation (7.3, above), centrifuge approx. 800-1000 rpm 10-15min.; drain supernatant

9.2. Re-suspend pellet in 0.5 ml ice-cold 30% ethanol

9.3. While slowly vortexing, add 4.5 ml ice-cold 0.15N NaCl with 0.1% Triton X-100 drop-wise

9.4. Separate 2.5 ml into two tubes for Endo+ and Endo-

9.5. Centrifuge 800-1000 rpm 10-15min.; drain supernatant

9.6. Re-suspend pellet in 35 mL Endonuclease Buffer

9.6.1. Add 1 mL Endonuclease III + Endonuclease Buffer (Endo III:Buffer = 1:10) to Endo+ sample

9.6.2. Add 1 mL of sterile glycerol to Endo- sample for control

9.7. Incubate in 37° water bath for 30 min. (up to 90min.)

9.8. When loading vials, do not add more Endonuclease Buffer, go to 10.2 of Loading section below

10. Loading

10.1.Re-suspend pellet in 35 mL Endonuclease Buffer (skip if sample has been treated with Endo III)

10.2.Add & re-suspend in 70 mL 2.25% melted (37°-RT) ULGT agarose

10.3.Pipette 100ml solution into warm-RT ½ dram vial pre-coated with 0.1% LGT

10.4.Place vials on ice for a minimum of 10min to allow gel to solidify
{ensure gel is solid & adhered!}

10.5.Store at 4° until alkaline treatment, cap vial if going to be held more than 48hr

11. Alkaline Treatment.

11.1.Cover gel with 1ml alkaline solution

11.1.1. Use pH 13.00 for all but extreme treatments – 20.0 min.

11.1.2. Use pH 12.15 for extreme treatments – 45.0 min.

11.2.Hold at room temperature in dark

12. Washing.

12.1.Remove alkaline solution & add 1 ml ice-cold TBS, then remove

12.2.Add 1 ml ice-cold TBS, hold on ice-4°C in dark for 10min. {neutralizes
remaining alkali}

12.3.Replace with 1 ml ice-cold TBS, hold on ice-4°C in dark for 20 min. minimum

12.4.Store covered with TBS at 4°C in dark until flow analysis, cap vial if going to be held longer than 48 hr

13. Flow Analysis.

13.1.Transfer vials to 65^oC heat block to melt agarose {approx. 45-60 sec, longer time increases variation}

13.2.Pipette 50 mL of melted gel/cell solution into 4ml polystyrene round-bottom tube

13.3.Add 100 mL ice-cold Solution A, gently mix w/ gel/cell solution & hold on ice

13.4.After 30 sec., add 300 mL ice-cold Solution B with AO (6.67 mg AO/ml Solution B)

13.5.Hold on ice for 3-30 min. (final AO concentration = 14.79mM/2X10⁵ to < 2X10⁶ cells)

13.6.Add 0.5-6ml chicken red blood cells (CRBC)

13.6.1. CRBC = standard to allow comparison of data obtained on different days

13.6.2. preferably 10% of cells < CRBC concentration < 30% of cells

13.6.3. volume of CRBC should not exceed approx. 10ml

13.7.Perform flow analysis (do not vortex)

2.25% ULGT agarose (SeaPrep agarose, Cambrex (FMC BioProducts)), store at 4°

1. Boil 50 ml PBS in a 50 ml Falcon tube

2. Add 1.125 g of Agarose slowly, allowing the gel to dissolve completely.

0.1% LGT stock (MetaPhor agarose Cambrex (FMC BioProducts)), store at 4°

1. Boil 50 ml ddH₂O in a 50 ml Falcon tube

2. Add 0.05 g of Agarose slowly, allowing the gel to dissolve completely.

Endonuclease buffer: solvent = dd H₂O, store at 4°

Compound	Concentration	Component for 50 ml
Tris base	20 mM	0.1573 g
NaCl	100 mM	0.292 g
500 mM EDTA	1 mM	100 mL

Adjust pH to 7.4

Alkaline solution: solvent = 0.15N(0.9%) NaCl, store at RT

Compound	For 50 ml of pH 13.00	For 50 ml of pH 12.15
0.2M NaOH	approx. 20ml	approx. 12ml
0.2M KCl	approx. 30ml	approx. 38ml
500 mM EDTA	5 ml/50 ml (50 mM final)	5 ml/50 ml (50 mM final)

Solution A*: solvent = glass-bottled water, store at 4°: pH 1.3

Compound	Concentration	Component for 50 ml	Component for 100 ml
Triton X-100	0.1%	50 mL Triton X	100 mL Triton X
HCl	0.08 N	1 ml of 4 N HCl	2 ml of 4 N HCl
NaCl	0.15 N	0.438 g NaCl	0.876 g NaCl

*pH 1.3

Solution B: solvent = glass-bottled water, store at 4°

Compound	Concentration	Component for 100 ml	Component for 200 ml
Na₂HPO₄	0.126 M	1.789 g Na₂HPO₄	3.598 g Na₂HPO₄
Citric acid*	0.037 M	0.777 g Citric Acid	1.554 g Citric Acid
NaCl	0.15 N	0.877 g NaCl	1.754 g NaCl
Na₂EDTA	1 mM	0.038 g Na₂EDTA	0.076 g Na₂EDTA

*pH 6 after sodium diphosphate + citric acid

Check off sheet for Alkaline Unwinding Protocol

Cells_____________

1. Cool on ice:

1.1. ______% EtOH (see #7, below)

1.2. 0.9% NaCl (no additives) pH 7.4 pH ______

1.3. 150mM Tris Buffered Saline (TBS) pH 7.5 pH ______

2. Coat ½ dram glass vials with 30 ml LGT 0.1% agarose.

3. Dry vials in 65^oC heat block & melt tube of 2.25% ULGT.

4. Harvest @10⁶ ±Rx cells.

5. Centrifuge @ 800 rpm for 10’ (____rpm/____’ actual).

6. Gently resuspend cells in 1 ml 37° DMEM+1.5mg/ml BSA [ ] (pH_____).

7. Hold on ice until ice-cold (<10 min.).

8. While vortexing ____, add 3 ml cold ____% EtOH dropwise (____% EtOH final).

9. Hold on ice minimum 10 min (____’ actual).

10. Centrifuge @ 800 rpm for 10’ (____rpm/____’ actual).

11. Gently resuspend in 0.5 ml cold ____% EtOH.

12. While vortexing ____, add 5 ml cold 0.15N NaCl.

13. Spin down @ 800 rpm for 10’ (____rpm/____’ actual) & DRAIN WELL.

14. Gently resuspend pellet in ____mL 4° Endonuclease Buffer (35 mL/sample).

15. Add ____mL 2.25% RT-37° ULGT agarose (70 mL/sample).

16. Pipette 100 mL cell/gel solution into RT-37° pre-coated vial:

17. Hold on ice for a minimum of 5’ for gel to solidify (____’ actual).

< Stopping Point>

Alkaline Treatment

18. Cover gel w/ 1ml alkaline solution (pH______), RT in dark 45’ (____’ actual).

19. Replace alkaline solution with 1 ml ice-cold TBS, hold on ice for 10’ (____’ actual).

20. Replace TBS with 1 ml ice-cold TBS, hold on ice for at least 20’ (____’ actual).

21. Drain TBS and store at 4°C until flow analysis.

<Stopping Point>

Flow Analysis

22. Transfer vials to 65^oC heat block to melt agarose (<1 min.).

23. Add approx. 3ml CRBC to tubes.

24. Gently add 50 mL of melted gel solution into 4ml polystyrene RB tube.

25. Add 100 mL ice-cold Solution A (pH 1.3 [ ]). Flick tube vigorously.

26. After 45 sec., add 300 mL ice-cold Solution B (pH6 [ ])+AO (6.67mg AO/ml B).

27. Perform flow analysis immediately, end < 3’ from AO addition.

Fluorometric DNA Unwinding Assay
(Macro assay)

Birnboim H.C. and Jevcak J.J. Fluorometric method for rapid detection of DNA strand breaks in human white blood cells produced by low doses of radiation. Cancer Research 41:1889-1892 1981

Baumstark-Khan C. Alkaline elution versus fluorescence analysis of DNA unwinding. Method in Enzymology 234:88-102 1994

Note: Error in published protocol – Use 0.1 M HCl and NOT 1 M HCl

1. Harvest cells and adjust to 5 x 10⁵/ml in FADU buffer.
Cell must have been fed the day prior to harvest

2. Keep 2-4 mls in reserve as B and T samples.

3. Treat P samples with g irradiation or H₂0_2.

4. Add 1 ml 0.1 M NaOH to each sample.

5. Add 1 ml of 0.1 M HCl to T samples only.

6. Follow protocols listed below for B, T and P samples..

7. Sonicate with the Branson Sonifier 250 in G. Martin lab
Duty Cycle set at Constant

Output control set at ~5 .
USE EAR PROTCTION!!!!!.

8. Use 15 ml polypropylene tubes.

B samples T samples P samples

(Background) (Total) (test)

1 ml in FADU buffer 1 ml in FADU buffer 1 ml treated sample in FADU buffer

1 ml 0.1 M NaOH 1 ml 0.1 M NaOH 1 ml 0.1 M NaOH

Sonicate 20 sec 1 ml 0.1 M HCl incubate for 30’ at 20°C

incubate for 30’ at 20°C incubate for 30’ at 20°C 1 ml 0.1 M HCl

1 ml 0.1 M HCl Sonicate 5 sec Sonicate 5 sec

Resonicate 5 sec Add 1 ml of Hoescht 33358 Add 1 ml of Hoescht 33358

Add 1 ml of Hoescht 33358 Read in fluoremeter Read in fluoremeter

Read in fluoremeter

FADU buffer 1 liter

Final concentration FW Amount added to 1 liter

5 mM Tris base 121.1 0.6 g

0.45 mM EDTA 372.24 0.167 g

135 mM NaCl 58.44 7.88 g

3 mM KCl 74.56 0.223 g

5 mM Na₂HPO₄ 141.96 0.71 g

1 mM KH₂PO₄ 136.09 0.13 g

pH to 7.4

If you are going to do a peroxide treatment add 10 ml of 0.01 M Ferric Chloride (FeCl₃) for every ml of FADU Buffer

Hydrogen Peroxide dilution

10 ml of 30% H₂0₂ to 10 mls of H₂0 for the stock solution.

Add 11.4 ml of the stock solution to 1 ml of cells for a 100 mM concentration

Sorensen’s Buffer

Na₂HPO₄ KH₂PO₄

(0.15 M/l: 10.6 g/ 500 ml H₂0) (0.15 M/l; 10.2 g/500 ml H₂0)

80 ml 14 ml

pH should be 7.6

Hoechst 33358 (Make fresh on the day of experiment)

Lab stock is 590 mg/ml

Add 11.31 ml/10 ml of Sorensen’s Buffer)

Use fluoremeter in C. Distesch’s lab.

Set excitation for 355

Set emission for 450

Use glass cuvettes and 2 ml of solution

Calculations

P-B = estimate of the double stranded DNA

F is the fraction of double stranded DNA

F=(P-B)/(T-B)

F_D / F_D=0

F_D=0 are DNA double strand fractions of untreated cells

SSF (strand scission factors) = -ln(F_D / F_D=0)

Micro alkaline unwinding with PicoGreen

Ref. Analytical Biochemistry 270:195-200 (1999)

Purpose_________________________________________________________________________________________________________________________________________

Before you begin -- make sure you can get time on the fluorometer and sign up.

Treatment and cells used:______________________________________________.

1. Check pH of 0.025 M NaOH.

a) Dilute 1 mls of 0.1 M NaOH with 7 mls of water for 0.05 M NaOH.

b) Add 250 ml of TE, 250 ml of lysing buffer and 2 ml of 0.025 M NaOH.

c) pH should be 12.15 ± 0.02.

d) Adjust pH of 0.025 M NaOH if pH is not correct.

2. Harvest cells and adjusted to 240,000 cells/ml in TE or 25 mg/ml of tissue in TE.

3. Add 25 ml of cells to microtiter plates. Plate each sample in quadruplicate.

4. Add 25 ml of lysing solution supplemented with PicoGreen 20 ml/ml of lysing solution.

5. Put plate on ice in the dark for 40 minutes

6. At this point you can turn on the fluorometer to make sure it is warmed up.

7. Add 200 ml of 0.025 M NaOH to all wells except Control + Water. Add 200 ml of water to those wells.

8. Read on the fluorometer at 485/530 at 5, 10, 15, 20, 25, and 30 min.

1. Lysing buffer 2. TE, 10 mM Tris 1 mM EDTA pH 7.4

9 M Urea

0.2 M EDTA 3. 0.1 M NaOH

0.1% SDS pH 10

Pancreas Parenchyma Dissociation
for CK Sort

Notes: This procedure produces material that is adequate for FISH, telomere FISH, and AP-PCR.

Keep all solutions and tissue on ice at all times; pre-cool in advance. Cool all syringes, needles and tubes on ice. Be sure to use nuclease-free water and BSA for all solutions.

Procedure

1. Place the pancreas parenchyma tissue (up to 5 mm³) into a 60 mm petri dish with 1 ml of cold PBA (1X PBS + 0.1% BSA) on ice.

2. Decant the PBA and mince the tissue in 1 ml of Inhibitor Mixture (PBA + 5 mM EDTA, 2 mg/ml Aprotinin). Mince into very small pieces (less than 0.5 mm if possible).

3. Incubate on ice for up to 10 minutes. Note, begin the 10-minute countdown at the start of mincing.

4. Using a cold 3 cc syringe w/LuerLok gently mix up and down the tissue several times to dissociate the cells. Add a cold 18-gauge needle to the syringe and continue mixing several more times into a cold sterile 17 x 100 mm polypropylene tube with cap. Insert the needle through the plastic cap to prevent splashing and loss of cells.

5. Finally, draw up the cell mix into the 3 cc syringe, lock a filter holder containing a 74-micron mesh to the syringe, filter the dissociated tissue through the 74-micron mesh into a pre-cooled 15 ml FCS-coated tube.

6. Rinse the petri dish with 1 ml of cold PBA + 5 mM CaCl₂, 5 mM MgCl₂ to collect any remaining cells, and filter the PBA/CaCl₂/MgCl₂ through the mesh into the same 15 ml tube. Gently mix the cells.

7. Centrifuge the tubes at 1700 rpm (500 x g) for 10 minutes at 4°C.

8. Decant the supernatant (carefully keeping the cell pellet intact), and add 500 ml of 1% paraformaldehyde/PBS, re-suspend the cells gently and incubate on ice for 20 minutes.

9. Add 1 ml of PBA to stop fixation. Gently mix the cells.

10. Centrifuge the tubes at 1700 rpm (500 x g) for 10 minutes at 4°C.

11. Add 500 ml of cold 0.1% Triton X-100/PBA directly to cells and incubate on ice for 3 minutes.

12. Add 1 ml of cold PBA to the Triton X-100/PBA. Gently mix the cells.

13. Centrifuge the tubes at 1700 rpm (500 x g) for 10 minutes at 4°C.

14. Prepare CK antibody mix (for each cell population of 1 million cells)

Add 6 ml of AE1/AE3-PE antibody
For non-a-CK: add 3 mouse-PE antibody

15. Decant supernatant and add 100 ml of PBA to the cells and re-suspend the cells. Transfer 10 ml of cells to a separate 15 ml FCS-coated tube and add 90 ml of PBA; label this tube as “negative antibodies”. Then add 14 ml CK antibody mix (or, 11 ml non-a-CK) directly to cells, gently re-suspend the cells, and incubate on ice for 60 minutes in the dark or overnight. If the cells pellet is large increase the amount of PBA to compensate. Also, adjust as necessary the amount of CK antibody.

16. Add 2 ml of cold PBA, and gently re-suspend the cells.

17. Centrifuge the tubes at 1700 rpm (500 xg) for 10 minutes at 4°C.

18. Decant and re-suspend cells in 300 ml of DAPI/10% DMSO. Adjust the amount of DAPI/DMSO as needed. Use solutions without NP-40 (the bottle with the GREEN label for increased signal strength).

19. Cells can be stored at –20°C until sorted.

Anti-Cytokeratin (CAM 5.2)-FITC labeled, Becton Dickinson Catalog #347653

Mouse IgG2a-FITC labeled, Becton Dickinson Catalog #349051

Anti-Rabbit IgG-RPE labeled (Fab)2, Sigma Catalog #P-8172

Anti-Cytokeratin (AE1/AE3) unlabelled, Roche Catalog #1-124-161;
AE1/AE3 antibody needs to be conjugated Custom PE conjugation by Intergen at 1-800-468-7436

For sorting use protocol SD1123/Nancy DAPI/PE/NC in CaCl₂.
Sort onto slides for FISH and into tubes for AP-PCR (minimum 1000 cells for AP-PCR).

Slide post-fixation for CK+ sorted pancreatic cells dropped from flow cytometer

Make up methacarn 3:1 methanol to acetic acid and put in freezer.
Mark location of the drop with a diamond pen and label slide in PENCIL.
Allow slides to air-dry over night after sort and before fixation.
Put slides in a Coplin jar or boat of ice-cold methacarn. Do NOT submerge the pence marks or else small bits of graphite will adhere to the cells and reduce the number of readable cells.
Let sit for 10 minutes in the hood.
Remove excess methacarn by blotting ends of slides into a paper towel.
Dispose of methacarn in appropriate bottle in the hood.
Rinse twice with 2x SSC for 2 minutes each (in the hood – it still smells).
Dehydrate in 70%, 85%, 90% and 100% ethanol for 3 minutes each.

Store the slides above the liquid nitrogen and note their location in the freezer log.

Two color Anti-CldUrd and
anti IdUrd Ab Staining

1. Cells have been pulsed with IdUrd and CldUrd

26 mg of IdUrd/10 ml of 0.05M Tris (pH 9.5) for a 10mM solution, filter sterilize.

5 mg of CldUrd/10 ml of PBS for a 10 mM solution, heat to 37 °C, filter sterilize.

2. Cells are frozen in 10% DMSO until the day of labeling.

3. Add 3 ml PBS to thawed cells and spin down, decant.

4. Add 1ml PBS + 5% NGS, spin down and decant.

5. Resuspend pellets to 150ml with PBS /5% NGS

6. Add 450 ml membrane shredding solution (100ml PBS, 500ml NONIDENT P-40 (IGEPAL CA-60), 20 mg EDTA).

7. Put samples on ice for 15 minutes, vortexing every 3 minutes.

8. Add 4N HCl 600ml and agitate for 30 minutes at room temperature.

9. Add 1ml 1M Tris (pH 8.5). Spin down and decant.

10. Wash one time with PBS + 5% NGS.

11. Add 50 ml Br3 1: 10 (from CalTag detects CldUr or BU1/75 from Harlan BioSciences) in diluted in PBS/5% NGS, 1 hr at RT.

12. Wash 1 time in PBS/NGS.

13. Add 100 ml goat anti-mouse PE 1:50 (or 100 ml of goat anti-rat PE if you used BU1/75 antibody) incubate 1 hr at RT.

14. Wash 1 time in PBS/NGS.

15. Incubate in 10% normal mouse serum to block free sites on Goat anti-mouse PE for 15 min at 37°C.

16. Wash 1 time in PBS/NGS.

17. Resuspend in 100ml of FITC conjugated anti-BrdU 1:25 (B44 from Becton Dickinson recognizes IdUrd) diluted in PBS/5% NGS/NMS.

18. Incubate with Ab in dark for 1 hour at 37°C.

19. Add 1 ml PBS/5% NGS; spin down cells and decant.

20. Add DAPI with chicken RBCs (666 ml CRBC in 20 mls) in PBS.

References

Aten, J.A., Bakker, P.J.M., Stap, J., Boxchman, G.A., and Veenhof, C.H.N. 1992 DNA double labelling with IdUrd and CldUrd for spatial and temporal analysis of cell proliferation and DNA replication. Histochemical J. 24:251-259.

Jaunin, F., Visser, A.E., Cmarko, D. Aten, J.A., and Fakan, S. 1998. A new immunocytochemical technique for ultrastructural analysis of DNA replication in proliferating cells after application of two halogenated deoxyuridines. 46(10):1203-1209.

White, R.A., Pollack, A., Terry, N.H.A., Meistrich, M.L., and Cao, S. 1994. Double labelling to obtain S phase subpopulations: application to determine cell kinetics of diploid cells in an aneuploid tumor. Cell Prolif. 27:123-137.

Lymphocyte Proliferation
Tritiated Thymidine

1. Prepare plate:

a. For each mouse there will be 9 wells prepared (3 wells with anti-CD3, 3 wells with ConA and 3 background wells).

b. Anti-CD3 wells: Calculated the total amount of anti-CD3 needed by multiplying # of wells by 50ml of this into the appropriate wells and incubate for 30 minutes at room temperature. Then tip plates and carefully aspirate off fluid in these wells. Gently wash wells once with 50ml of PBS.

c. Prepare growth media. Calculate the total mls of growth media needed by multiplying the total numbers of wells by 200ml = X. Take X and divide it by 14.8 to get a fraction to multiply all components of growth media by

Growth Media

RPMI 13.0ml

Fetal Calf Serum 1.5ml

Pen/Strep 0.15ml

2-Mercaptoethanol 0.15ml

Example: To calculate growth media for 6 wells:

6 wells X 200ml = 1200ml = 1.200ml

1.200ml = 0.081 (multiplication factor)

14.8 ml

RPMI 13.0ml X 0.081 = 1.053ml

FCS 1.5ml X 0.081 = 0.122ml

P/S 0.15ml X 0.081 = 0.012ml

2 ME 0.15ml X 0.081 = 0.012ml

d. Make media with ConA

Calculate total amount of media with ConA needed by multiplying # of wells by 200ml. Divide this number by 80 to get the number of mls of ConA needed to make a 1:80 dilution of ConA in growth media.

e. Fill wells with 190ml of appropriate media. Fill surrounding wells with

200ml of PBS.

2. Place 250,000 cells per well (12.5ml if 2.0 X 10⁶cells/100ml).

3. Incubate the plate at 37^o in a 5% CO₂ incubator.

4. After 56 hours add 1 mCi of tritiated thymidine to each well (20 ml of dilution) and return plate to incubator. [Dilute tritiated thymidine stock (20 Curies/mmol) to 1 mCi/20 ml (1 ml stock + 19 ml media)]

5. At 72 hours harvest cells or place entire plate into the freezer until the plate can

be harvested.

6. Harvesting cells: (harvester is in hood in T-152)

a. Label vials and place on harvester rack.

b. Turn the vacuum pump on and flip the switch to harvest.

c. Place a filter on the harvester (rough side up) and wet filter with water (push button on side of hand held aspirator 3 times quickly) and watch to see water coming out the other end.

d. Place plate in harvester and aspirate out material from the wells and rinse wells by pushing button 8 times. Watch to make sure all water is gone from all wells.

e. Dry the filter by aspirating absolute methanol through and allow the filter to dry for a minute or two.

f. Flip switch to raise and raise the bar. Check to see that all filter discs are suctioned to the top of the harvester mesh. Place the rack under it, lower the bar, flip the switch to harvest and raise and lower bar 5 or 6 times to shake filter discs loose. Check to see that all discs are in the vials.

g. Wash off filter area with ethanol before harvesting another set.

h. When done, rinse harvester and associated tubing with bleach solution followed by copious amounts of water.

i. Turn the vacuum pump off.

7. Add 2 mls of BetaMax to each vial and cap them. Fill 3 vials with 2 mils of BetaMax (do in room with scintillation counter) only to be used as background counts.

8. Load the scintillation counter as follows:

a. Place rack of standards (C-14, H-3, and BKG) on the back right side of the counter. The samples standards should be on the left side of the rack as they are counted from left to right.

b. In a separate rack, load the 3 background tube on the left hand side of the rack with program card #2 in it. Leave one space empty after the background tubes and then load consecutively the samples to be counted in triplicate. Should a sample not be done in triplicate, the samples should be loaded leaving the spaces empty when there aren’t three samples. The halt rack (red colored rack) should be placed in after the last rack of samples. The racks should be loaded from the back to the front such that the standards are in the very back and the halt rack is nearest to you.

c. Using the menu screen, check program #2 by selecting “Review and Edit User Program.” Move the cursor to “2 H3-Kathy” and check specifications. They should read as follows:

Counting time = 1.00 minutes

Scintillator = Liquid

Isotope 1 = ³H

Isotope 2 = None

If any of these are incorrect, enter the correct information. If you “get lost” in the program you can hit “Previous Menu” until you get back to the main menu and begin again.

d. When you wish to begin, select “Automatic Counting” and then “Start.” Also sign in the record book on the right hand side of the counter can be followed. As samples are counted a printout is made on the printer to the right of the counter. When the samples are done counting, they should be removed and the scintillant placed in a brown bottle appropriately labeled for disposal.

9. Analysis of data:

Scintillation counter will calculate Mean and standard deviations for each set of triplicates. Scan data to see if there are spurious data points that should be omitted. For each mean, subtract the mean background counts from the mean stimulated counts. This data can either be compared directly or expressed as percent of control.

Materials: RPMI 1640 INVITROGEN/GIBCO Labs Cat. # 430-1800EB

FCS Hyclone Labs

                                                ConA

Pen/Strep SIGMA Cat. # P-3539

2-ME Aldrich Cat. # M370-1

Anti-CD3 mAb BD (Pharmingen) Cat. # 01081D

98 wells plates Nunc Cat. # 1 63320

PHA VWR Cat. # DF0528-56

³H 20 curies/mmol

Drug treatment @ 37 ^oC for the optimal time interval.

TUNEL

Original protocol from Kevin Henne (David Hockenbery lab FHCRC)--with some modifications

1. Fix cells (in suspension or adhered to slide) in 10% formalin overnight (minimum 16 hrs).

2. For cytospins: cytocentrifuge cells onto silane-prep slides at 1600 rpm for 4 mins, air dry. (Can store slides at 4°C for future use.)

3. Rinse in 1X PBS for 5 mins.

3a. Strip nuclear proteins using proteinase K (stock 20 mg/ml ) 1:300 dilution in PBS. Incubate 15 mins @ RT. (Can be omitted if cellular destruction occurs. Do NOT use for cytospins.)

3b. Rinse 4 times in 1X PBS. (1 minute per rinse)

4. Quench endogenous peroxidases: immerse slide in 2% H₂O₂ for 10 mins. [3.4 mls 30% H₂O₂ in 50 mls PBS] Repeat second time in 2% H₂O₂. Rinse well 4 times in 1X PBS, 1 minute per rinse. (H₂O₂ may inhibit TdT buffer.)

5. Cover slides with 1X TdT buffer (30 mM Trizma base pH 7.2, 140 mM Na Cacodylate, 1 mM Cobalt chloride) for 5 min @ RT. Cover cytospins with 100 ml of primary enzyme solution; incubate @ RT for 2 hrs.

Specimens + positive control: 1.2 ml TdT enzyme + 0.75 ml biotin-labeled dUTPper 100 ml 1X TdT buffer

Negative control: 0.75 ml biotin-labeled dUTP per 100 ml 1X TdT buffer

[Prepare Vectastain ABC solution: 1 drop A + 2.5 mls PBS. Vortex. Add 1 drop B. Vortex. Incubate on ice at least 30 mins.]

6. Rinse 4 times with 1X PBS.

7. Block nonspecific avidin binding with 2% BSA in PBS for 10 min @ RT. Rinse 4 times in PBS.

8. Cover cytospins with Vectastain ABC prepared at least 30 mins earlier. Incubate @ RT for 30 mins. Rinse 4 times in 1X PBS.

9. Prepare substrate immediately prior to use: 2 mls DAB (0.5 mg/ml) + 250 ml 8% NiCl + 1 ml H₂O₂. Incubate from 3 to 10 mins @ RT. Stop reaction with ddH₂O. At this point, can check under microscope. Can leave in ddH₂O and coverslip later.

10. Counterstain with 0.25% methyl green for 10-15 mins. Rinse in ddH₂O.

11. Dehydrate and cleanse with xylene:

95% ETOH-------1 rinse (40 dips)

100% ETOH------2 rinses (40 dips each)

xylene-------------3 rinses (5 mins total)

[Make sure to change xylene regularly, approximately after every 2 runs]

12. Coverslip using Permount.

--------------------------------------------------------------------------------------------

**Preparation of TUNEL positive control: Camptothecin treated CEMs**

1. Put 0.5 mls of CEMs in 15 ml conical tube.

2. Add 5 mls RPMI + 10% FBS containing 3 mM camptothecin (3 ml 10 mM camptothecin stock in 10 mls media).

3. Incubate 1 hr @ 37°C.

4. Pellet cells. Resuspend in 2 ml RPMI + 10% FBS without camptothecin.

5. Transfer cells to 30 mm tissue culture dish and incubate at least 18 h.

6. Pellet cells and resuspend in 10% formalin solution.

--------------------------------------------------------------------------------------------

TUNEL reagents

Stocks:

1. 300 mM Tris pH 7.2 (10X stock) store at 4°C

18.17 g Tris in 500 ml dH₂O

2. 1.4 M Cacodylic Acid (10X stock) store at 4°C

8.96 g Cacodylic Acid in 40 ml dH₂O

[Use gloves and mask]

3. 10 mM Cobalt Chloride (10X stock) store at 4°C

.095 g Cobalt Chloride in 40 ml dH₂O

[Use gloves and mask]

4. 40 mM MgCl₂ (10X stock) store at 4°C

.325 g MgCl₂ in 40 ml dH₂O

5. 1 mM Dithiothreitol (10X stock) store at 4°C

6.17 mg DTT in 40 ml dH₂O

6. 0.1 M Acetic Acid store at room temp.

3 ml glacial acetic acid in 500 ml dH₂O

7. 0.1 M Sodium Acetate store at room temp.

4.1 g Sodium Acetate in 500 ml dH₂0

8. 3.0 M NaCl (10X stock) store at room temp.

17.53 g NaCl in 100 ml dH₂O

9. 300 mM Sodium Citrate (10X stock) store at room temp.

8.82 g Sodium Citrate in 100 ml dH₂O

Solutions:

1. DN buffer (store at -20°C)

30 mM Tris pH 7.2 [ 5 ml 10X Tris stock]

140 mM Cacodylic Acid [ 5 ml 10X Cacodylic Acid stock]

4 mM MgCl₂ [ 5 ml 10X MgCl₂ stock]

0.1 mM DTT [ 5 ml 10X DTT stock]

**Add 30 ml dH₂O to make 50 ml buffer

2. TdT buffer (store at -20°C)

30 mM Tris pH 7.2 [ 5 ml 10X Tris stock]

140 mM Cacodylic Acid [ 5 ml 10X Cacodylic Acid stock]

1 mM Cobalt Chloride [ 5 ml 10X Cobalt Chloride stock]

**Add 35 ml dH₂O to make 50 ml buffer

3. TB buffer (store at room temp.)

300 mM NaCl [10 ml 10X NaCl stock]

30 mM Sodium Citrate [10 ml 10X Sodium Citrate stock]

**Add 80 ml dH₂O to make 100 ml buffer

4. 2% BSA (store at 4°C)

0.8 g BSA in 40 ml dH₂O

5. 8% Nickel Chloride (Make fresh weekly) store in dark at 4°C

0.25 g in 3.1 ml dH₂O

6. 2.5% Methyl Green (store at room temp.)

183.75 ml of 0.1 M Acetic Acid

66.25 ml of 0.1 M Sodium Acetate

5 g Methyl Green

**Filter before each use if not used daily

7. DAB (carcinogenic)

-Add 20 ml 0.05M Tris pH 7 into vial containing 10 mg DAB powder
using hypodermic syringe

Freeze 2 ml aliquots in capped test tubes. Store up to one month.

Dispose of DAB by bleaching overnight

Product Qty Company Product # Price

Standard Vecta Stain

Elite ABC Kit 1 Vector laborites PK-6100 $150

3,3’-Diaminobenzidine

tetrahydrochloride (DAB) 1 Polysciences Inc. 04008

SECTION 16 Lab Forms

List all chemicals that come into the lab

Chemical Name Amt. Location Date Initials

________________________________________ _______ ___________ ________ ________

Record of media supplies to other labs

Charge is 15 min. of flow time for 1 liter of media

Lab Group Type of media Amount Date of prep. Date of Charge

____________________________ _______________ ________ ________________ ________________

____________________________ _______________ ________ ________________ ________________

____________________________ _______________ ________ ________________ ________________

BioRad Confocal Photography Record

Stains used:_______________________________________________________

_______________________________________________________

Slide ID: ________________________________________________________

________________________________________________________

Photo ID __ __ __ __ __ __ __ __ Slide ID:________________

Z step Yes No _____(nM)

PMT1 PMT2 PMT3 Mixer PMT1 PMT2 PMT3

Save Mixer A Iris A

Mixer B Gain B

Mixer C Bkg L C

Emm.Filter

Laser (nm) ________ Laser power________ Magnification_______ Kalman _______

Description:____________________________________________________________________

______________________________________________________________________________

Photo ID __ __ __ __ __ __ __ __ Slide ID:________________

Z step Yes No _____(nM)

PMT1 PMT2 PMT3 Mixer PMT1 PMT2 PMT3

Save Mixer A Iris A

Mixer B Gain B

Mixer C Bkg L C

Emm.Filter

Laser (nm) ________ Laser power________ Magnification_______ Kalman _______

Description:__________________________________________________________________

____________________________________________________________________________

Photo ID __ __ __ __ __ __ __ __ Slide ID:________________

Z step Yes No _____(nM)

PMT1 PMT2 PMT3 Mixer PMT1 PMT2 PMT3

Save Mixer A Iris A

Mixer B Gain B

Mixer C Bkg L C

Emm.Filter

Laser (nm) ________ Laser power________ Magnification_______ Kalman _______

Description:__________________________________________________________________

____________________________________________________________________________

Continuation Page _______ Date:____________

Photo ID __ __ __ __ __ __ __ __ Slide ID:________________

Z step Yes No _____(nM)

PMT1 PMT2 PMT3 Mixer PMT1 PMT2 PMT3

Save Mixer A Iris A

Mixer B Gain B

Mixer C Bkg L C

Emm.Filter

Laser (nm) ________ Laser power________ Magnification_______ Kalman _______

Description:__________________________________________________________________

____________________________________________________________________________

Photo ID __ __ __ __ __ __ __ __ Slide ID:________________

Z step Yes No _____(nM)

PMT1 PMT2 PMT3 Mixer PMT1 PMT2 PMT3

Save Mixer A Iris A

Mixer B Gain B

Mixer C Bkg L C

Emm.Filter

Laser (nm) ________ Laser power________ Magnification_______ Kalman _______

Description:__________________________________________________________________

____________________________________________________________________________

Photo ID __ __ __ __ __ __ __ __ Slide ID:________________

Z step Yes No _____(nM)

PMT1 PMT2 PMT3 Mixer PMT1 PMT2 PMT3

Save Mixer A Iris A

Mixer B Gain B

Mixer C Bkg L C

Emm.Filter

Laser (nm) ________ Laser power________ Magnification_______ Kalman _______

Description:__________________________________________________________________

____________________________________________________________________________

Lieca Confocal Record Sheet

Date___________

Experiment Description: _______________________________________

_________________________________________________________

Argon Laser ____________ Power_______________

Kryton Laser____________ Power_______________

HeNe Laser______________

UV Laser _______________

File Name:________________________

Photo ID	PMT 1	Off set 1	PMT 2	Off Set 2	PMT 3	Off set 3

Dye Name	Preference	Base pair specificity	Exc. max. (nm)	Excitation sources	Emm. max. (nm)	Comments
Acridine Orange	RNA DNA	No No	460 502	Mercury arc lamp Argon 457, 488 nm	650 526	DNA/RNA Discrimination
7-AAD	DNA(RNA)	Weakly GC	545	Mercury arc lamp HeNe543 nm	647	Weakly permeant; live-dead discrimination
DAPI	DNA	AT	359	Mercury arc lamp Argon 360 nm	461	Ploidy analysis and cell cycle studies; minor groove binding
Ethidium Bromide	dsDNA(RNA)	No	510	(Mercury arc lamp) Argon 488 nm	595	Impermeant; base pair Intercalator
Hoechst 33342	dsDNA	AT	346	Mercury arc lamp Argon 360 nm	460	Permeant; cell cycle studies; minor groove binding
Hoechst 33258	dsDNA	AT	346	Mercury arc lamp Argon 360 nm HeNe 543 nm	460	Impermeant; cell cycle studies; minor groove binding
Propidium Iodide	DNA(RNA)	No	536	Argon 488 nm	617	Impermeant; live-dead discrimination
Pyronin Y	DNA RNA	No No	559 560	(Argon 488 nm; Mercury arc lamp) HeNe 543	569 573	Permeant; DNA/RNA discrimination with Hoechst33342
SYTOX^TM Green	DNA(RNA)	No	504	Argon (457) 488 nm	523	Impermeant; cell cycle studies live/dead discrimination
TO-PRO^TM-1	DNA(RNA)	No	515	Argon 488, 514 (Mercury arc lamp)	531	Impermeant; cell cycle studies
TO-PRO®-3	DNA(RNA)	No	642	HeNe 633; Diode 635 (Mercury arc lamp)	661	Impermeant; cell cycle studies
TO-TO®-1	DNA(RNA)	No	514	Argon 488, 514 (Mercury arc lamp)	533	Impermeant; cell cycle studies
TO-TO®-3	DNA(RNA)	No	642	HeNe 633; Diode 635 (Mercury arc lamp)	660	Impermeant; cell cycle studies
YO-PRO^TM-1	DNA(RNA)	No	491	Argon 457, 488 (Mercury arc lamp)	509	Impermeant; cell cycle studies
YO-YO®-1	DNA(RNA)	No	491	Argon 457, 488 (Mercury arc lamp)	509	Impermeant; cell cycle studies

Pull the Fire Alarm or call 9-911

Call 206.543-0467

Lymphocyte Preparation from Whole Blood

Section 6 Protocols for FISH and Antibody Stains

General Guidelines for BioRad (scanning confocal microscope in the Immunology Cell Analysis Facility)

Preparation of cDNA from total RNA (Reverse Transcription, RT)

ENZO RNA prep for generation of cRNA

AFFYmetrix RNA prep for generation of cRNA using IVT kit

Section 11 Macros, SAM, GeneTraffic hints

Lymphocyte Preparation from
Whole Blood