MLS 450 - City University of New York



I. PURPOSE:

To describe aseptic techniques used for handling sterile products and/or mammalian cells

II. REFERENCES:

Freshney, R.I. Culture of Animal Cells (1983) Alan r. Liss, Inc., New York

III. DOCUMENTATION REQUIREMENTS

Certification of hood

IV. DEFINITIONS:

V. EQUIPMENT AND MATERIALS:

A. Equipment

1. Class 100 biological safety cabinet (Laminar flow hood)

2. Pipet-Aid

3. 20-200 μl micropipetters

B. Materials

1. 70% ethanol in a squirt bottle (CAUTION! THE STOCK BOTTLE OF ETHANOL SHOULD NOT CONTAIN ANY METHANOL)

95% ethanol 367 ml

Distilled H20 132 ml

2. Pipets, sterile, of appropriate size.

3. Biohazard waste container (also known as red bags)

4. Latex or nitrile surgical type gloves

5. Clean liquid waste container (a good thing to use is empty sterile water bottles)

6. Paper towels

VI. PROCEDURE:

A. Personal hygiene

1. Hair should be tied back

2. Beards should be covered with a face mask

3. Talking at the hood should be minimized. Do not talk at all if possible.

4. A mask will assist in prevention of coughing if this might be a problem

5. Always spray gloves with 70% ethanol before putting them into the hood

B. Clothing

1. Clean lab coat.

2. Close fitting powder-free surgical gloves made of latex or other non-latex material are required.

C. Preparation of hood working area

1. Fan. It is best to have the fan running continuously but when there are long periods of down time, such as weekends, the fan can be turned off. After restarting, run the fan for at least 30 min before using.

2. Working surface. Spray gloves with 70% ethanol and then spray the entire work surface and wipe up with a paper towel.

3. All equipment such as pipetmen should be washed with 70% ethanol before being brought into the hood.

D. Pipetting

1. Transfer pipettes. Use sterile-wrapped, cotton-plugged transfer pipettes with a Pipet-aid. Open the first few inches of the package at the cotton-plugged end and attach the pipette to the Pipet-aid before removing the rest of the package.

2. Pipette tips for P-10, P-20, P-200, and P-1000 must be sterilized by autoclaving. They should be removed from the autoclave while still warm and brought to the laminar flow hood for drying so as not to draw dirty air into the box. The cool, dry, sterile pipettes can be stored outside of hood in their tightly closed boxes when not in use. The autoclave tape, indicating their autoclave history should be left on the box and can be use to seal the box.

E. Changing tissue culture medium

1. Clean all reagent bottles with paper towels saturated with 70% ethanol before bringing them into the hood.

2. Bring flasks to be changed from the incubator into the hood.

3. Wash gloves with 70% ethanol

4. Open package of new flasks in the hood and remove new flasks to working surface. Seal the opened flask bag containing any unused flasks with tape to prevent it popping open outside of the hood.

5. Change the medium using appropriate protocol

6. Any drips or spills should be swabbed up immediately with ethanol.

7. If centrifugation is used, discard supernatant into the waste container by inverting the centrifuge tube. BE SURE NOT TO TOUCH THE TUBE TO THE WASTE CONTAINER.

8. Discard used pipettes and flasks into biohazard waste bags

9. Replace cells in the incubator

10. Remove media bottles and place parafilm around the lid to prevent entry of fungal spores before replacing them in cold room.

11. Add bleach to the waste bottle to a concentration of at least 10% and let stand for 30 min. Discard down the drain of the sink and rinse the bottle.

12. Wash down the surface of the hood.

13. Remove gloves and wash hands.

Proper media is crucial for the growth of cells and tissues in culture. While the actual medium used may be dictated by such variables as cell type, available equipment, experimental protocol, etc., all media must provide for the physical and nutritional requirements of the cells. The physical requirements include temperature, pH, osmotic pressure, etc. The nutritional needs vary with cell type but generally include: carbohydrates, amino acids, nucleic acids, lipids, vitamins, hormones, ions, and trace elements. Most media, especially the early formulations, attempted to mimic the in vivo milieu of the cells. To achieve this, complex natural body fluids such as plasma (actually serum is most often used) are included in the medium. Totally synthetic media (defined media) of known composition are becoming more common as the specific growth-promoting factors in the plasma (serum) are identified. Many of the media components are either heat or light sensitive (or both) and will deteriorate rapidly. All media should be refrigerated (but not frozen!) to reduce this lability. As a general rule media should be used within one moth of preparation.

OBJECTIVES:

1. To demonstrate the procedures for making and sterilizing media as well as doing checks for microbial contamination of media.

2. To provide media for subsequent labs.

REAGENTS:

The following stock solutions will be available in sterile form and should be handled using aseptic technique:

1. Minimum Essential Medium (MEM)

2. Rosewall Park Memorial Institute medium # 1640 (RPMI 1640)

3. Fetal Bovine Serum (FBS)

4. Non-essential amino acids (NEAA)

5. Vitamin mixture (V)

6. Penicillin/streptomycin (pen/strep)

7. Gentamicin

8. Fungizone

SUPPLIES:

1. 100 ml graduate cylinder

1. 10 ml sterile individually wrapped pipets

1. 20 – 200 ul micropipetters and pipet tips

PROCEDURE:

1. Practice aseptic technique in preparing both media. You will sterile filter the media after it is made but you should take care not to contaminate the reagents.

2. MEM Medium: measure the following amounts of each reagent (they do not have to be added in the order below) and mix in a 100 ml graduate cylinder:

Reagent amount

MEM 88 ml

FBS 10 ml

NEAA 1.0 ml

vitamin mixture 1.0 ml

pen/strep 0.1 ml

gentamicin 0.1 ml

fungizone 0.2 ml

3. RPMI Medium: to a second 100 ml graduate add:

reagent amount

RPMI 90 ml

FBS 10 ml

pen/strep 0.1 ml

gentamicin 0.1 ml

fungizone 0.2 ml

4. Pour the media from the first into the upper chamber of a 0.22 u Millipore filter unit (at this point the filter units are top-heavy and very easily tipped over - especially during the next steps).

6. Attach the vacuum hose to the side arm (this will be separated wrapped and must be attached aseptically to the bottom chamber) of the filter unit so that the cotton plug is retained.

7. Apply vacuum until all of the media passes into the lower chamber.

8. Remove the upper chamber and attach the lid provided.

9. Repeat steps 4 - 7 for the RPMI 1640 media.

10. Aseptically remove 2 drops (a 1 ml pipette works well) from each media bottle and add one drop to a separate blood agar plate and the second drop to a separate nutrient broth tube. Be sure to label each with your group number. Check for growth of contaminating microorganisms in the blood agar plates and nutrient broths at the start of the next lab.

11. Label each media bottle with media, name, group, and date.

12. Store your media in the tissue culture refrigerator at about 4o. For each laboratory your media should be warmed to room temperature or 37o if possible.

QUANTIFYING CELLS

In many studies an accurate measure of cell number or other quantifiable characteristic is required. The simplest of these measures is to count cells or to determine the total cellular protein concentration. In this lab three methods will be demonstrated. Manual and automated methods of cell counting will be compared. In addition total cellular protein will be measured. Each method has advantages and disadvantages. Manual cell counting using a hemacytometer is most often employed when there are few samples to count and when cell viability must be determined. Automated cell counting, on the other hand, is valuable when many samples must be counted and where cell viability is not critical. Because cells within a homogeneous culture vary in size the amount of protein per cell will also vary. The total cellular protein will be an average of the amount of protein in the cultured cells. It is often used when other quantitative cellular parameters are measured. For example enzymatic activity is normally related to total cellular protein content.

OBJECTIVES:

To demonstrate three techniques of quantifying cells. One or more of these methods may be used for routine maintenance of cells in future labs.

CELLS:

EG.7 cells are murine thymoma cells that grow in suspension

PROCEDURE:

1. E.G7 cells will be provided in a T-75 flask.

2. These cells grow in suspension and can be harvested easily as follows:

a. Gently swirl the flask to resuspend all cells.

b. remove the cap and pour the suspended cells into a 50 ml centrifuge tube.

3. Spin cells down in the bench-top centrifuge for 5 minutes at ¾ speed.

4. Pour off the supernatant (into disinfectant).

5. Wash cells by resuspending in 10 ml of HBSS.

6. Centrifuge as above.

7. Repeat the was twice more using 10 ml of HBSS.

8. After the final wash resuspend the cell pellet in exactly 1.0 ml of HBSS.

9. Remove 10 μl aliqouts in triplicate and add to three wells (use adjacent wells in columns 4, 5, and 6) of a microtiter plate (There will be a single plate for all groups to share) for protein assay described below.

10. Remove a 100 μl aliquot and mix (pipet up and down several times with the micropipettor) with 100 μl of trypan blue for cell counting using the hemacytometer as described below.

11. Remove a second 100 μl aliquot and add to exactly 10.0 ml of isoton solution for counting using the Coulter Counter as described below.

PROTEIN ASSAY:

1. Group #1 (group #4 in section 002) will set up a standard 1:2 serial dilution protein curve as follows:

a. Select a 96 well microtiter plate (with flat bottom wells)

b. Add 30 μl of distilled water to wells B through H of column #1

c. Remove 30 μl from the 2 mg protein standard and add to well 1 A in column #1

d. Remove 30 μl from the 2 mg protein standard and add to well 1 B ,mix well then remove 30 μl and add to well 1 C

e. Repeat steps c & d until all dilutions have been made.

f. Remove 30 μl from well 1 H and discard

2. Group #2 (group #5 in section 002) will set up triplicate standards to the as

follows:

a. Transfer 10 μl aliqouts from well 1 A to wells 2 A and 3 A

b. Do the same for wells B - H

c. This should set up triplicate values (in columns 1, 2, and 3) for each dilution of the standard

3. Group #3 (group #6 in section 002) will mix the protein reagents as follows:

a. Calculate the total volume of protein reagent needed

i. All standard and unknown well must be counted

ii. All wells will require 150 (l of reagent

iii. The total volume should be 1.0 ml greater than that required for all wells

iv. Total (in ml) = wells x 0.15 + 1.0 ml

b. Mix the reagents to make up the final reagent in the following ratio

i. Reagent A = 25 parts

ii, Reagent B = 24 parts

iii. Reagent C = 1 part

4. Add the protein reagent (150 (l/well) after the unknown and standard wells have been set up

5. Cover the plate with its lid

6. Transfer the plate to the incubator for 1 hour

7. Read absorbance using the ELISA Reader with a wavelength of 560 nm.

MANUAL MEASUREMENT OF CELL NUMBERS AND VIABILITY:

1. Clean a hemacytometer by wiping the surface with alcohol

2. After the alcohol dries place a coverslip on the hemacytometer

3. Using a micropipettor set at 100 μl, resuspend the cell:trypan mixture by pipetting up & down several times.

4. Add a small drop of mixture to the groove of the hemacytometer until the chamber fills completely.

5. Place the hemacytometer under the microscope at low power and locate the grid (refer to the diagram on page 13)

6. All grids have the same size and volume but the center grid is easiest to use

7. Count all of the clear cells in the entire center grid and record the number

8. Count all of the blue cells in the same grid and record the number

9. See page 14 for calculations of cell number and viability.

AUTOMATED MEASUREMENT OF CELL NUMBERS

1. Label Coulter Counter vials with the information for each cell type

2. Add 10.0 ml of Isoton solution to each counting vial

3. Gently resuspend each cell type by swirling the tubes

4. Using the micropipettor transfer 100 μl of each cell suspension to the corresponding counting vials

5. Count the vial three times using the Coulter Counter set at 0.5 ml

6. Record each count and use the mean

7. Calculate total cell counts and compare to the other measures

CELL COUNTING AND VIABILITY

The hemacytometer should be cover-slipped before adding a drop of the cell suspension/trypan blue mixture to the groove at the edge. The hemacytometer is divided into 9 major grids as described on the next page. Each of these grids is subdivided into smaller and smaller grids as shown. The area of any of the 9 major grids is 1 mm2 (1 mm x 1 mm). The height between the grid and the cover slip is 0.1 mm. Therefore the total volume of 1 major grid is 0.1 mm3 or 0.1 ul. This is equal to 10-4 ml.

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CALCULATING CELL VIABILITY

Viable cells are those that do not take up the dye and therefore should appear clear. Nonviable cells will be blue because damage to the membrane allows entry of the trypan dye. The viability can be determined when cells are counted using the hemacytometer. The % viability is calculated as the total clear cells divided by the total of both clear and blue cells times 100.

% viable = [clear cells/clear + blue] x 100

CALCULATING TOTAL CELL NUMBER

1. Hemacytometer Method:

Record the total number of clear cells in one of the major grids. Multiply this by 2 to correct for the dilution when trypan blue was added. This value is the number of cells in 10-4 ml. Therefore multiply this number by 104 to give the total cells per ml of your cell suspension and times the total ml of your suspension if you have more than 1 ml.

total cell number = counts x 2 x 104 x ml of suspension

2. Coulter Counter

Determine the average cell counts from three measurements. This represents the number of cells in 0.5 ml of the cells suspended in the isoton. Determine the dilution factor to be used to calculate total cell numbers.

PASSING CELLS

Very often cells cannot be maintained for long periods of time in individual culture containers (flasks). It is desirous to transfer cells to new flasks for a number of reasons: to grow more cells, to set up different experimental protocols, cells may die if left in a flask, or to alter cellular metabolism. This process of transfer from flask to flask(s) is known as "passing" and each time it is done the cells are said to be at 1 higher passage number. To be passed cells must first be harvested (removed) from the flask in which they are growing. This process can be done by physical methods (e.g. shake off or scrape off) or chemical methods (e.g. enzyme action or cation removal). The method of choice will depend upon the cells grown and specific experimental conditions, but usually should be as gentle as possible to limit cell damage and death. In passing cells a number of parameters must be considered including: plating efficiency, split ratio, minimum and maximum inoculum, growth rates, contact inhibition, attachment strength, and cell fragility. Throughout the first half of this semester you will harvest and/or pass various cell types by a number of different methods. However, by far the most widely used technique is the trypsinization/EDTA method described below.

FREEZING CELLS

An integral and extremely important aspect of tissue culture is to maintain frozen stocks of all cells that are grown. Frozen cell stocks are necessary to: maintain availability of early passage numbers of cells, prevent total loss of cell lines (due to contamination, equipment malfunction, personal error, etc., compare early and late passage cells, etc. Cells must be harvested and frozen by methods that cause the least cell damage. This usually requires the prevention of swelling when ice crystals are formed because the cell plasma membranes may be susceptible to rupture caused by the swelling. A number of procedures have been utilized to minimize this swelling. These include addition of agents such as glycerol, dimethylsulfoxide (DMSO), sucrose, serum, etc. The rate of temperature change has also been found to affect cell death, a very slow rate of cooling being the best. In this lab you will use freezing medium (normal growth medium for each cell type) containing 10% DMSO and a step-wise freezing protocol for two different cell types.

OBJECTIVES:

1. To demonstrate the techniques of harvesting, cell counting and viability testing, and passing cells.

2. To freeze both cell types in the event that either becomes contaminated or dies in the future.

CELLS:

1. Normal mouse skin fibroblasts [named MSF-1] - early passage cells isolated from mouse skin in a previous class. These cells grow to confluency and remain as contact inhibited cultures.

2. Murine B 16 melanoma - cells originally isolated from a spontaneous melanoma found on a black C57Bl/6 mouse in the 1950's and maintained in culture or in vivo. These cells are not contact inhibited but will die if allowed to remain confluent for more than 1 day.

PROCEDURE:

1. Follow aseptic technique using sterile supplies for both cell types.

2. Remove the medium from the flask of cells provided to each lab group by carefully pouring (do not allow the disinfectant to splash back into flask) into a beaker of disinfectant (wescodyne).

3. Add 5 ml of Hank's Balanced Salt Solution (HBSS) without calcium & magnesium to the cells, rinse gently, and pour off.

4. Repeat step #3 twice.

5. Add 1 ml of trypsin-EDTA solution and leave on the cells until they begin to float off. Do not allow trypsin to remain on the cells beyond this time!

6. (use the same sterile 5 ml pipette for steps 6 & 7) add 4 ml of complete medium (MEM for B16 melanoma cells & RPMI 1640 for mouse fibroblasts) to each flask and gently suspend cells by squirting liquid onto growing surface.

7. Transfer the medium with the cells to a 15 ml sterile centrifuge tube.

8. Spin the cell suspension at 1000 rpm (approximately 1/2 power) for 5 minutes.

9. Pour off the supernatant and carefully resuspend the cell pellet in 1.0 ml of HBSS (without calcium and magnesium) using a 1 ml pipette. Be sure that the pellet is completely broken up and that the cells are uniformly suspended.

10. Using a sterile micropipettor remove 100 ul aliquots from each cell suspension and add to 100 ul of trypan blue dye which you already placed in a small test tube (this tube will only be used to count cells and does not have to be sterile).

11. Count the cells and determine the cell viability using a hemacytometer according to the protocol outlined in Parts IV - VI (on pages 12-13).

12. Passing cells at 1:10 split ratio (high density): remove an aliquot (100 ul), which contains 10% of the total cells, and transfer to a new flask already containing 5 ml of the appropriate growth medium. NOTE: the following information should be written on each flask - date, medium, cell type and your group number.

13. Low density passage: remove a small aliquot (10 - 20 ul or about 1 drop) and put into a new flask with 5.0 ml of the appropriate medium.

14. Freezing cells: spin down the remaining cells at 1000 rpm. Resuspend the cell pellet in 1.0 ml of the appropriate complete medium containing 10% DMSO (this will be provided and labeled "freezing medium.")

15. Transfer the medium with cells to freezing tubes and label as follows: cell type, cell number, date, and group number. The tubes will be incubated for you at room temperature for 15 minutes, at 4o for 1 hour, at minus 5o degrees for 1 hour, then in the ultralow freezer (Revco) at - 60o.

MAINTAINING CELLS

To provide fresh nutrients, remove toxic metabolites, maintain pH, etc. the culture medium must be changed often. The frequency of these changes (feeding cells) will generally depend upon the individual cell types which are grown in culture and the cell number. Ideally, most cells are fed 3 times a week, some require feeding every other day or even every day, while others can survive with 2 feedings per week. Both of the cell types you have should be fed at least twice per week. Feeding simply requires the removal of the medium and replacement with fresh medium. If cells are near or at confluency then you must decide if they should be passed rather than fed.

Your cells will not survive over the weekend without feeding. You must feed (or pass) your cells on Friday and again next Tuesday. You will maintain these cells until the midterm exam. They can be maintained on a Tuesday/Friday schedule of feeding and or passing. Before you feed your cells you should examine them for contamination. If your cells are contaminated seal the cap tightly and put the flask into the orange autoclave bag. Do not leave contaminated cells in the incubator!

THAWING CELLS:

At some point in operations of most tissue culture labs frozen cell stocks must be brought out to replenish cultured cells for a variety of reasons already discussed. Like the procedure for freezing cells, this must be done in a manner, which minimizes cell damage and contamination. Experience has shown that the greatest cell viability is obtained when cells are thawed rapidly. This is directly opposite to the procedure for freezing cells. The viability of cells that are thawed will be determined by the freezing procedure, the conditions in which the frozen cells are kept, and the thawing procedure itself.

OBJECTIVE:

To demonstrate the technique of thawing and establishing cultures from frozen cells.

CELLS:

Both mouse skin fibroblasts and B 16 melanoma cells frozen in the previous lab will be utilized.

PROCEDURE:

1. Remove ampules from the ultralow freezer (or liquid nitrogen) and immediately place in a water bath at 37o. DO NOT immerse the tubes totally but be certain that the water level is below the bottom of the tube cap. Agitate by swirling the tube until the ice has completely melted (usually between 1 and 2 minutes). caution: if the ampule was stored in liquid nitrogen and the cap not sealed tightly the ampule may have filled with liquid nitrogen. This should not damage the cells but can be dangerous if the tube is sealed tightly before thawing. The rapidly boiling liquid nitrogen will cause enough pressure to build up in the tube that it will crack (with explosive force).

2. As soon as the ice has melted remove the ampule from the bath and wipe thoroughly with an alcohol wipe. This may remove the labels on the tubes so be sure that the information is recorded beforehand.

3. All subsequent operations should be carried out using aseptic technique.

4. Transfer the contents of each tube to separate sterile 15 ml centrifuge tubes using a sterile pipette.

5. Add 5-10 ml of the appropriate growth medium, mix thoroughly, and spin down cells at 1,000 rpm for 5 minutes.

6. Pour off the supernatant and carefully resuspend the cell pellet in 1.0 ml of HBSS without calcium and magnesium. Be sure that the pellet is completely broken up and the cells are uniformly suspended.

7. Take 100 ul aliquots from each cell suspension and add to 100 ul of trypan blue dye in small test tubes.

8. Measure and calculate the % cell viability and the total cell number.

9. Set up one new flask for each cell type and examine them in 2 - 3 days for attachment and growth. Record your findings and report the observations in your lab report. These flasks can be discarded if you have other flasks that contain viable, uncontaminated cells growing.

10. You will be responsible for maintaining each cell type in culture until the midterm exam when they should be turned in. You may want to freeze down some of each now or at the next passage.

11. If your cells are contaminated or do not grow then you should attempt to obtain viable cells from other groups when they are passed.

SERUM REQUIREMENTS FOR GROWTH

Optimal conditions for cell growth will depend upon a number of factors that have already been discussed in connection with culture media composition. In most tissue culture labs, and in this course, media containing serum is routinely used for maintaining cultured cells. Both the species and concentration of the serum may affect cell growth. These factors as well as the total cost (sterile fetal bovine serum which is free of mycoplasma and tested for pyrogens, toxins, etc. is expensive) must be considered. In this lab the optimal concentrations of fetal bovine serum for growth of both cell types will be determined. For the purpose of this lab the optimal conditions will be those (FBS %) which give the greatest number of cells. The protocol used to determine these conditions can, in general, be used to measure the optimal concentration of any factors which might affect cell growth.

OBJECTIVES:

1. To determine the concentration of fetal bovine serum which gives the best growth of each cell type.

2. To demonstrate the general protocol which can be used to obtain the optimal concentration of factors that affect cell growth.

CELLS:

1. Mouse skin fibroblasts which were obtained and passed in the previous lab.

2. B16 melanoma cells which were passed in the previous lab.

PROCEDURE:

1. Set up the following sterile 15 ml tubes:

tube # medium (ml) FBS (ml) % FBS

1 10.0 0.0 0.0

2 9.75 0.25 2.5

3 9.5 0.5 5.0

4 9.0 1.0 10.0

5 8.5 1.5 15.0

6 8.0 2.0 20.0

2. One set of these tubes should be set up for B16 melanoma cells and one set for fibroblasts. The medium used above should be the appropriate complete medium for each cell type except that no FBS has been added yet.

3. Harvest the cells of each type from one flask that you have been maintaining. Be sure that the flask is near confluency so that you will have sufficient cells to set up this experiment. If you have any doubts consult with your instructor. Use the trypsinization protocol described in a previous lab to harvest cells.

4. Check cell viability and calculate total viable cells as previously described.

5. Transfer 5 x 104 viable cells (for both MSF-1 and B16 melanoma cells) to each of 6 separate T-25 flasks (DO NOT ADD THE CELLS TO THE TUBES OF MEDIA MADE ABOVE!).

6. Add 5.0 ml of the appropriate medium from each of the 6 tubes to the flasks and label each flask to designate the % serum and cell type.

7. These cells must be fed with the appropriate % serum media again on Monday before the next lab.

8. Check the cells daily for growth and if any reach confluency note the days since plating that this occurs.

9. After 7 days (at the beginning of the next lab) cell numbers should be measured as follows:

a. remove the medium and wash cells 3 x 5 ml of HBSS without Calcium and Magnesium. Add 1.0 ml of trypsin/EDTA to each flask. Because these cells will not be maintained, aseptic technique is not necessary.

b. when cells begin to come off the flask wash off the remaining cells using the trypsin/EDTA previously added to each flask with a pipette and uniformly suspend them (in each flask).

c. remove 100 ul aliquots from each flask and transfer to counting vials that contain 10.0 ml of isoton.

d. determine the total number of cells by counting in the Coulter Counter with the setting on 0.5 ml.

e. calculate the total cell number for each flask then plot total cells/flask versus serum %.

Cultured cells provide an ideal source for studying metabolic processes. The use of radiolabelled precursors is the most common method for studying both anabolic and catabolic pathways. Potential loss of radiolabel compounds from the desired anatomical site or metabolism in other organs are not issues when cultured cells are used. The potential hazards of radioactive substance use to whole organisms are also eliminated in cultures. Cholesterol synthesis is a normal anabolic pathway that can be measured using a radiolabelled precursor. Most commonly sodium acetate containing either 3H or 14C tags is added to cells for 1 hour to 2 days. After the incubation cells are harvested and the cholesterol extracted. The amount of radioactivity in the cholesterol extract will indicate the rate of the synthetic pathway.

Another method for analyzing metabolic pathways is to measure enzymatic activity. Most pathways are controlled by a "rate-limiting enzymatic step." The enzyme that catalyzes this step may be called the rate-limiting enzyme. For cholesterol synthesis the rate-limiting enzyme is called 3-hydroxy-3-methylglutaryl Coenzyme a reductase (HMG Co reductase). The activity of this enzyme is correlated with the total synthetic rate of cholesterol. The activity of this enzyme is easy to measure in cultured cell homogenates. An advantage of enzymatic assays is the ability to directly measure inhibitors of activity. Early studies in culture cells identified some inhibitors of cholesterol metabolism that led to development of drugs (lipitor and others that are among the biggest selling drugs on the market) that reduce serum cholesterol in humans.

In this lab we will measure the activity of some very common enzymes in our cell cultures. Our ability to measure enzymes in intact whole cells may be limited by the location of the enzyme within the cell and/or the permeability of the cells to the substrate. In most cases the cells must be broken up to free cytoplasmic enzymes or to isolate membrane-bound enzymes. Among the easiest methods of cell homogenization is the hand-operated homogenizer such as the Dounce homogenizer. This is simple and fast. It disrupts cell membranes by creating shear forces that tear the cells apart.

After homogenization it is often desirable to prepare crude cell subfractions to partially isolate the desired enzyme. The simplest cell fractionation involves centrifugation. A low speed spin will pellet cell debris and nuclear material while other fractions remain in the supernatant. A second moderate speed spin can be used to spin down membrane-containing components. This is called a microsome fraction. The supernatant of this spin will contain free cytoplasmic components. If this isolation is used the fraction containing the target enzyme can be analyzed.

OBJECTIVES:

1. To demonstrate how to harvest cells rapidly by.

2. To demonstrate a simple homogenization technique.

3. The measure activity of two different cellular enzymes.

CELLS:

1. Mouse skin fibroblasts from previous labs.

2. B16 melanoma cells from previous labs.

PROCEDURE:

1. These cells will not be passed so aseptic technique is not required.

2. Wash the cells three times with ~5 ml of HBSS as described previously.

3. Add 10 ml of HBSS and scrape the entire surface of the flask (Be careful not to spill any!).

4. Using a 10-ml pipet, wash the flask surface with the cell suspension. This will also serve to break up cell clumps.

5. With the same pipet, collect the cell suspension and transfer it to a 15-ml centrifuge tube.

6. Spin down the cells and discard the supernatant.

7. Resuspend the pellet in 1.0 ml of PBS.

8. Transfer the entire 1.0 ml suspension to a homogenizer.

9. Homogenize the cells as follows:

a. Do not force the pestle down rapidly! This might cause the liquid and cell suspension to spray out of the homogenizer.

b. Push the pestle to the bottom of the tube.

c. The upward stroke can be done rapidly.

d. Homogenize the cells by using a total of 30 strokes (up and down = 1 stroke)

10. Remove 50 (l and determine cell viability (you should not even see cells).

11. Remove triplicate 10 (l aliqouts for protein assay as described previously (be sure

to use columns 1-3 for the standard curve and columns 4-6 for you cellular protein

samples.

12. Remove triplicate 50 (l aliqouts for measurement of alkaline phosphatase activity.

These aliquots should go in wells in columns 7-9 of the same microtiter plate.

13. Remove triplicate 50 (l aliqouts for measurement of lactate dehydrogenase (LDH)

activity. These aliquots should go in wells in columns 10-12 of the same microtiter

plate.

14. After all samples have been added to the wells, the reagents should be added:

a. Protein reagent should be added to all wells that contain samples or standards in columns 1 - 6. Be sure to run triplicate reagent blanks in wells 4 - 6 H. 150 (l/well of the reagent is required.

b. Alkaline phosphatase reagent should be added to all wells that contain samples in columns 7 - 9. Be sure to run triplicate reagent blanks in wells 7 - 9 H. 150 (l/well of the reagent is required.

c. LDH reagent should be added to all wells that contain samples in columns 10 -.12. Be sure to run triplicate reagent blanks in wells 10 - 12 H. 150 (l/well of the reagent is required.

15. Place the plate in the incubator for 30 minutes.

16. Read the plate once each using the following wavelengths.

a. Protein 560 nm

b. Alkaline Phosphatase 405 nm

c. LDH 650 nm

17. Return the plate to the incubator and leave for 1 hour before taking a second reading.

Cells may often be difficult or impossible to grow even in the richest possible medium. To grow these cells it may be necessary to add special ingredients and/or growth factors. Growth factors are hormone-like molecules produced by certain cells of an organism which act to stimulate or to sustain growth of other cells of the organism. Examples of growth factors include fibroblast growth factor (FGF), nerve growth factor (NGF), interleukins (IL-1,2,3, etc., and platelet-derived growth factor (PDGF). In this lab the growth of lymphoid cells which require the growth factor IL-2 will be examined. IL-2 is in fact quantified by a tissue culture assay (using cells similar to those used in this lab and serial dilutions of IL-2) to define the unit of activity.

Measurement of cell growth by counting cells in a hemacytometer or by Coulter Counter is difficult to do when multiple samples must be measured. The simplest method to measure cell growth is to analyze DNA synthesis. This is typically done by a radioactive precursor assay. 3Thymidine, a precursor for DNA, is added to the cultures for a given period of time (usually form 1 - 24 hours). After incubation, the cells are harvested and the amount of radioactive thymidine that was incorporated into DNA is determined. Dead cells will incorporate no thymidine and faster growing cells will incorporate more thymidine than slower growing cells. In recent years the use of radioactive precursors has been limited because of problems of storage of the radioactive waste generated. New, non-radioactive assays have been developed cell growth and viability.

The MTT dye assay is a spectrophotometric assay of cell viability. MTT is a small organic compound 3-[4,5 dimethylthiazol-2-y]-2,5 diphenyltetrazolium bromide that can enter all cells. In the cytoplasm of cells the MTT is broken down by a mitochondrial enzyme into an insoluble black precipitate. Dead cells do not have the enzyme and can not form the black precipitate. Thus, the amount of precipitate that is formed is directly proportional to the number of viable cells in the culture. To quantify cell viability the precipitate can be dissolved in a detergent and the absorbance measured in a spectrophotometer. The absorbance in the unknown cultures can be compared to absorbances of cells grown in a standard curve of IL-2 concentrations. In this manner the concentration of IL-2 in an unknown can be determined.

OBJECTIVES:

1. To gain experience in handling cells which grow in suspension.

2. To identify and determine the concentration of growth factor in an unknown solution.

CELLS:

HT2 - a murine lymphoma cell line which absolutely requires the presence of IL-2 to grow

REAGENTS:

1. HT2 media: complete media containing all nutrients needed for growth but without IL-2.

2. IL-2 stock solution: A stock solution of IL-2 sufficient to make up a serial dilution standard curve starting at 1000 U/ml.

3. Unknown media A, B, & C: normal growth media containing different units of IL-2.

PROCEDURE:

1. Make a Il-2 standard serial dilution curve:

a. Use the top 2 rows (A & B) of a microtiter plate.

b. Add 50 (l of growth medium to wells A2 - A12 and B2 - B12.

c. Add 100 (l of the IL-2 stock (2000 U/ml) to wells A1 & B1.

d. Transfer 50 (l from A1 & B1 to A2 & B2.

e. Mix well with the micropipet then transfer 50 (l to the next set of wells.

f. Continue the serial dilution until all wells of rows A & B have been completed.

G. Discard 50 (l from wells A & B 12.

2. Add 50 ul of your unknown to duplicate wells of the plate in rows D and E.

3. Harvest HT2 cells by gentle agitation then spin down cells.

4. Wash cells 3 x 10 ml of sterile HBSS (with Ca/Mg)

5. Resuspend in 10 ml of complete medium and determine cell viability and total cell number.

6. Dilute or resuspend cells to give a final concentration of 2 x 105 cells/ml.

7. Add 50 ul of cell suspension to all standard and unknown wells to give 10,000 cells/well.

8. Allow cells to grow overnight in the incubator.

9. In the morning add 10 ul of MTT dye (5 mg/ml) to each well.

10. Six hours later add 33 ul of 10% SDS + .02 N HCl to each well.

11. Incubate overnight in the incubator.

12. Measure absorbance at 650 nm in the ELISA reader.

13. Calculate the concentration of IL-2 in your unknown sample.

A typical cell of a viable culture growing in log phase will eventually divide to give two daughter cells. These, in turn, will also divide giving rise to other cells. The interval between the appearance of a cell and the generation of two new daughter cells from it is known as the cell cycle. The total length of time needed to complete one full cycle (the cell cycle traverse) varies with the type of cell and also the growth conditions. Transformed (i.e. cancer) cells often have shorter cell cycles. The cell cycle has been subdivided into four distinct phases:

M - the actual physical separation of two cells from one

G1 - the time interval between M and S phases

S - the time when DNA replication occurs

G2 - the time interval between S and M phases

For most cells G1 is the longest phase followed by S, G2, and M respectively. If a single cell is isolated its cell cycle phase can be determined by measuring the DNA content. G1 = single copy of all DNA, G2 = double copy of all DNA, and S = DNA content intermediate between single and double copies. Individual cell DNA content can be measured with a fluorescent dye that binds specifically to DNA [Not to protein or RNA!] and a flow cytometer or cell sorter (see figure 12.14 in your ). A number of fluorescent dyes can be used for this purpose. In this lab we will use propidium iodide. This is a convenient dye because it can be excited by visible light [488 nm which is the wavelength of an argon laser] and it fluoresces strongly in the orange-red visible region. If unsynchronized cells are growing in log phase then the pattern should resemble that shown in figure 12.15 of your .

If all cells of a culture could be collected in the same phase then these cells are called synchronized. The fluorescent pattern would then show only one peak corresponding to the amount of fluorescence of that specific phase. Unfortunately the cell cycle traverse varies considerably from cell to cell and thus cells which are synchronized will lose that synchrony within 1 - 2 cell cycles.

Cells can be synchronized by a number of methods including actual separation of cells according to phase as well as the accumulation of cells in a single phase due to the addition of drugs (blockers) which prevent cells from continuing through the cycle.

If normal log phase cells are incubated in the presence of a blocker for longer than the cell cycle traverse then all cells should be in the phase blocked by the drug. When the blocker is washed out all of the cells will resume the cell cycle traverse at the same time and will therefore be synchronized.

OBJECTIVES:

1. To demonstrate the normal procedure for obtaining synchronized cells using three different cell cycle blockers.

2. To determine the phase that cells are in when treated with unknown blockers.

CELLS:

1. Murine EG.7 Thymoma cells

2. B16 melanoma also from your cultures

CAUTION:

Cell cycle blockers such as colchicine, cycloheximide, and cytocholasin B are highly toxic. Cytocholasin B is especially dangerous since it may be a carcinogen and it is dissolved in dimethyl sulfoxide (DMSO). DMSO is a solvent that can penetrate the skin bringing solutes with it into the body.

PROCEDURE:

Each lab group must determine the phase that cells are in when treated with an unknown cell cycle blocker. However all groups must use cells from the same source to reduce differences between samples and control cultures. In this lab we will set up only 4 flasks of B16 melanoma and 4 flasks of MSF-1 cells as follows:

1. Groups #1 & 4 will harvest 1 near-confluent flask of B16 melanoma cells by the normal trypsinization procedure.

2. Groups #2 & 5 will harvest 1 near-confluent flask of normal mouse fibroblast cells by the normal trypsinization procedure.

3. Each group will determine cell counts and viability normally.

4. Groups #3 & 6 will set up 4 new T-25 flasks for B16 melanoma and 4 for MSF-1 cells. 5 ml of normal growth medium should be added to each flask. Label the flasks 1 – 4.

5. Groups #1 & 4 will remove 4 aliquots from their B16 melanoma cell suspension so that each contains 5 x 105 viable cells and add them to each flask.

6. Groups #2 & 5 will remove 4 aliquots from their MSF-1 cell suspension so that each contains 5 x 105 viable cells and add them to each flask.

7. Groups #3 & 6 will then add the blockers as follows:

flask # addition volume

1 Control [no blocker 10 ul

2 unknown for Groups # 1& 4 10 ul

3 unknown for Groups #2 & 5 10 ul

4 unknown for Groups #3 & 6 10 ul

8. After at least 24 hours, harvest cells by the normal trypsinization protocol.

9. Spin down the cells and pour off the supernatant.

10. Buzz the pellet on a vortex.

11. While vortexing slowly add 2.0 ml of ice-cold 70% ethanol (in distilled water) to fix the cells.

12. If cell can not be stained and assayed immediately, they should be left in ethanol until ready.

13. PI can bind to double stranded RNA as well as DNA. To prevent this from happening the fixed cells will be treated with RNase to remove any RNA prior to staining.

14. Cells will be stained with propidium iodide just prior to running the flow cytometer.

15. Each group will get the print out from the flow cytometer analysis for all groups. You should determine what phase your cells are in from the data.

Many animal tumor cells are sometimes maintained by transplanting to suitable host animals. There are a number of reasons why this is routinely done:

1. Cells may be difficult to grow in culture.

2. Cells may in some way be altered by long periods of growth in culture. This might be simple or more complex changes associated with adaptation to the in vitro environment or can also be actual genetic changes that might be accelerated by the culture conditions.

3. Passage of transformed cells back to the natural host will at least demonstrate if they are still tumorigenic.

4. Growing cells in vivo can provide a much greater number of cells, usually at a much cheaper price.

5. Individual experiments involving the host's response to tumor growth can be examined.

Normally cells are transplanted back into the original (syngeneic) strain. A number of factors must be considered when transplanting tumor cells to animals including:

1. Site of injection of cells

2. Volume of cells injected (may be site dependent)

3. Number of cells injected, may depend upon:

a. minimum inoculum - MTD (minimum tumoricidal dose)

b. rate of growth

c. host factors and experimental parameters

Human tumor cells may also be passaged in animals under the proper circumstances. If human cancer cells are transplanted to normal animals they will not grow because the host will mount an immune response against the "foreign" human cells. "Heterotransplants" can, however, succeed if the host is immunocompromized. This is most commonly done by immunosuppressing mice or by using a genetically immunoincompetent strain such as the nude mouse.

In this course the in vivo passage of cells will require 2 lab sections. Part I will involve the injection of cells into the host and part II the isolation of viable cells from tumors when they have attained an appropriate size (because this is difficult to control the date of part II is subject to change). The procedure used in part II will be the same used to isolate primary cultures from spontaneous tumors.

PART I: INOCULATION OF ANIMALS

OBJECTIVES:

1. To demonstrate the procedure for isolation and injection of cells into an animal host.

2. To obtain animals which have tumors suitable for isolation of tumor cells.

ANIMALS:

The C57Bl/6 strain of black mouse from which the tumors cells originally came.

CELLS:

Bl6 melanoma cells.

PROCEDURE:

1. Harvest Bl6 melanoma cells from 1 confluent T-75 flask by flotation in cation-free medium as follows:

a. wash cells 3 x 10 ml with HBSS without calcium/magnesium.

b. Incubate cells with 10 ml of HBSS without calcium and magnesium for 15 minutes in the incubator or until cells float off from flask.

c. when cells are off remove the cells and spin them down at 1000 rpm for 5 minutes.

2. Resuspend cells in 10.0 ml of HBSS without calcium and magnesium.

3. Remove 100 ul aliquot and determine cell viability and total cell number by trypan blue dye exclusion.

4. Spin down cells and resuspend in appropriate volume of HBSS so that dilutions can be made to give the following:

Mouse # Cells injected volume of injection ear tag

1 5 x 103 50 ul -

2 5 x 104 50 ul 1 L

3 5 x 105 50 ul 1 R

4 5 x 106 50 ul 1 R&l

5. Examine mice weekly for appearance of tumors.

ISOLATION OF PRIMARY MURINE FIBROBLASTS

Primary explants, or cells, are those which come directly from fresh tissue. Originally, all tissue culture was done on primary cells but this had the disadvantage that only short term growth of cells was achievable (before modern tissue culture techniques were developed).

Isolation of specific cell types for long-term culture from fresh tissue can be difficult. A number of factors must be considered when establishing primary cultures. First, sterility and aseptic technique are extremely important, though often difficult to achieve, because fresh tissues may already contain potentially contaminating microorganisms. Next, the method for tissue disruption must be chosen to increase cell recovery but at the same time reduce cell damage, death, and contamination with other cell types. Finally, methods must be utilized which will insure that the desired cell type is isolated and grown.

OBJECTIVES:

1. To demonstrate the technique and difficulties of establishing primary cultures from fresh tissue.

2. To generate primary cultures of murine skin fibroblasts.

ANIMALS:

C57Bl/6 mouse

PROCEDURE:

1. Sacrifice the mouse by cervical dislocation after anesthesia.

2. Shave the back with a razor.

3. Disinfect the skin with 70% ethanol.

4. Carefully remove the skin from the back and cut away as much fat as possible. Be careful not to puncture the peritoneal cavity.

5. Remove two sections (about 1 cm2) of the skin and place into two petri dishes.

6. Using sterile forceps carefully press a sterile glass coverslip onto one of the sections of skin.

7. Gently add 10.0 ml of complete medium (RPMI + 10% fetal bovine serum) without dislodging the coverslip.

8. These cells must be fed normally being careful not to dislodge the coverslip.

9. To the second petri dish add 5 ml of HBSS and gently tease with forceps for 1 minute then remove the HBSS.

10. Add 2 ml of enzyme cocktail, mince the skin with sterile scissors, then incubate at 37o in the incubator.

11. Continue to occasionally mince the tissue.

12. At the end of the laboratory section transfer the petri dish to the refrigerator and incubate overnight.

13. Add 10 ml of RPMI + 10% fetal bovine serum to the dish to counteract the enzymes.

14. Harvest the medium with a pipette and transfer to a 15 ml centrifuge tube.

15. Spin cells down at 400 x g for 5 minutes.

16. Wash 2 x 5.0 ml with complete medium (RPMI + 10% FBS).

17. Remove an aliquot from the final wash to determine cell number and viability.

18. Set up a T-25 flask with 5 ml of complete medium.

19. Transfer the cells to the flask.

20. Feed these cells with fresh medium once before the next laboratory.

Monoclonal antibody technology was developed by Kohler and Milstein in the early 1970's. A monoclonal antibody can be thought of as a purified antibody that recognizes a specific antigen. The monoclonal antibody can be generated in very large quantities. This methodology has revolutionized not only immunology but almost every other area of research. It has already had a tremendous impact on our ability to analyze biological samples for small quantities of many substances and will continue to have an increasing role in many assays. To understand and appreciate the power of this technique a brief description of the background for this technology is necessary.

BACKGROUND:

Antibodies are generated and secreted by B lymphocytes in the body. A single B lymphocyte will not make several different antibodies (i.e. those which react with several different antigens) but rather will synthesize only a single antibody which will react with a single antigen. It is usually not possible to isolate any specific antibody from the serum because of the very large number of different antibody molecules present in the blood. The only way to obtain pure antibodies (monoclonal antibodies) would be to isolate the specific B lymphocytes that secrete the proper antibody. Unfortunately even though we may be able to isolate a single lymphocyte (by clonal dilution), these cells do not grow in culture. Kohler and Milstein found a way around this problem and were able to grow clonal B lymphocytes in culture that synthesized and secreted monoclonal antibodies of predefined specificity.

HYBRIDOMA METHODOLOGY: (see figure 8-1)

To illustrate this technique the example which Kohler and Milstein first used will be followed, i.e. monoclonal antibody formation against sheep red blood cells (SRBC). A suitable animal, the mouse, was chosen and immunized several times to SRBC. Production of antibodies against the SRBC was monitored by normal serological techniques. When the mouse had high titers of antibodies against SRBC the spleen (a source of high numbers of B lymphocytes) was removed and lymphocytes isolated. These cells were fused to murine myeloma cells (transformed cells of the B lymphocyte lineage which are immortal in culture) to give hybrids of B lymphocyte/myeloma. Many of the hybrids will retain the growth and immortality characteristics of the myeloma parent cells and also the antibody production of the B lymphocyte parent.

Once formed these hybrids must be separated from the original myeloma cells because the fusion is less (probably < 1% efficient) than total and the unfused myelomas normally grow better than the hybrids. This selection process can be accomplished if the original myeloma cell line used for the fusion is a specific mutant. The mutant myeloma cells chosen are those which lack an enzyme called hypoxanthine guanine phosphoribosyl transferase (HGPRT). This enzyme catalyzes the formation of purine nucleosides (precursors of DNA synthesis) from ribose and either hypoxanthine or guanine. This pathway (see figure 8-2) is called the salvage pathway for purine nucleoside synthesis. Purine nucleosides are normally synthesized by the pathway shown in figure 8-3 from simple precursors. Even though myeloma cells lack HGPRT they can grow because they

[pic]

figure 8-1

utilize the normal pathway. Selection of the hybrids is done by preventing growth of the mutant myeloma cells by incubating the fused cells in a selection media called HAT MEDIUM. HAT medium contains all necessary nutrients for growth but also has: hypoxanthine, aminopterin, and thymidine. Aminopterin is an analog of a coenzyme (f-THFA) necessary for the normal pathway (de novo synthesis) of purine nucleoside synthesis to occur (see figure 8-3). In the presence of aminopterin the de novo synthesis is blocked because it competes for binding to the enzymes with the f-THFA. Thus cells are forced to use the salvage pathway to obtain the needed purine nucleosides. Because the myeloma cells lack the HGPRT enzyme of the salvage pathway they can not synthesize purines nucleosides by either pathway when aminopterin is added and, therefore, will not be able to grow. Splenocytes (mostly lymphocytes) won't grow in culture under any conditions. Hybrids formed by the fusion of a splenocyte to a myeloma cell will be the only cells that can grow as the HGPRT enzyme will be obtained from the splenocyte parent cell. These hybrids will utilize the hypoxanthine as a precursor of purine nucleosides. HAT medium thus allows us to isolate only the lymphocyte:myeloma hybrids from all other parent cells or other possible hybrids.

Selection is usually done at low concentration of cells in many (10 - 20) 96-well tissue culture plates. If the proper hybrids are formed then small colonies of cells will eventually appear in the wells which contain those hybrids. These hybrids must be screened for the production of antibodies (in our case against SRBC). If antibodies are formed they will be secreted into the medium so that we can test for their presence by analyzing the medium of each well. Since there may be a large number of wells (20 plates x 96 wells/plate) this is usually done with a simple assay such as the ELISA.

Once we have identified the wells positive for antibodies against SRBC those hybrids must be collected (individually) and tested to be certain that only a single antibody (from a pure hybrid) is present. The best method to do this is to clone each positive colony. Cloning can be accomplished by limiting dilution and replating in large number of 96-well plates. When colonies grow up from this step they are then re-screened for antibody production. Most labs will reclone hybrids once or twice more to be certain that the hybrid is pure. A pure hybrid will secrete a pure (monoclonal) antibody that recognizes a specific antigen only. The monoclonal antibody (obtained from the medium of the hybrid (hybridoma) is then tested for specificity, isotype, and characterized. The hybridoma is maintained in culture to collect the antibody secreted into the medium. Often to collect large quantities of the monoclonal antibody the hybridoma cells are injected into the peritoneal cavity of syngeneic mice where they will grow (as ascites tumors) and secrete the monoclonal antibodies into the peritoneal fluid (ascites fluid). Collection of the ascites fluid will yield high concentrations of the monoclonal antibody.

[pic]

[pic]

[pic]

figure 8-2

[pic]

figure 8-3

OBJECTIVES:

1. To learn the basic methods utilized to isolated splenocytes from immunized mice.

2. To fuse splenocytes to myeloma cells.

3. To attempt to select hybridomas which may produce monoclonal antibodies to the immunogens.

ANIMALS:

C57Bl/6 mice which have previously been immunized to B16 melanoma (one of the mice injected with B16 cells to grow tumors).

CELLS:

P3K - a mutant murine myeloma cell line which lacks the HGPRT enzyme.

REAGENTS:

1. Fusion medium - normal growth medium (RPMI 1640 + 10% FBS) containing 30% polyethylene glycol (PEG).

2. HAT medium - normal growth medium containing hypoxanthine, aminopterin, and thymidine.

PROCEDURE:

1. Sacrifice the immunized mouse by cervical dislocation after anesthesia.

2. Soak the mouse briefly with 70% ethanol.

3. Place the mouse on a dissecting table with its left side facing up.

4. Make a shallow incision in the left flank with scissors and move the skin away from the area.

5. Cut into the peritoneal cavity taking care not to hit any organs, especially the intestines.

6. Grasp the spleen with forceps and lift it slightly to gently tease away the surrounding fat and tissue.

7. Place the spleen between two sterile glass slides (keep them directly above the petri dish) and gently squeeze until the membrane has ruptured.

8. Rinse the slides into the petri dish with normal growth medium.

9. Transfer the cells to a 50 ml centrifuge tube.

10. Harvest the P3K cells (they grow in suspension) by shaking gently then pouring the cells into a 50 ml centrifuge tube.

11. Spin down both cell types at 400 x g for 5 minutes.

12. Discard the supernatants and add 5.0 ml of cold 0.87 % ammonium chloride to the splenocytes and let stand on ice for 10 minutes.

13. During this time, determine the total number of viable P3K cells available by resuspending the pellet in 50 ml of PBS and removing a suitable aliquot for measuring viability and counting cells.

14. After the 10 minute incubation add 5 ml of growth medium to the splenocytes and spin down as above.

15. Resuspend the pellet in 25.0 ml of growth medium.

16. Determine the total number of viable splenocytes by removing a suitable aliquot from this sample and counting normally.

17. Calculate the number of myeloma cells needed to give a 1:4 ratio of myeloma to spleen cells.

18. Pool the proper number of myeloma and spleen cells (1:4) in a single 15 ml centrifuge tube and spin down cells.

19. Pour off the supernatant and tap the tube gently to break up the pellet.

20. Add 0.5 ml of fusion medium and centrifuge immediately at 400 x g for 4 minutes. Do not pour off supernatant. Time is critical! No more than 8 minutes should elapse before next step.

21. Add 5.0 ml of growth medium. Do not resuspend the pellet.

22. Spin down at 400 x g for 5 minutes.

23. Pour off supernatant and resuspend the pellet in 25 ml of normal growth medium (RPMI 1640 + 10% FBS and antibiotics) (not HAT medium!).

24. Transfer to a T-25 flask and allow to stand upright in the incubator overnight.

25. On the following day spin the cells down and resuspend in 25 ml of HAT medium.

26. Transfer the cell suspension to 3-4 96-well microtiter plates by addition of 0.1 ml (use the repeat pipettor) to every well except the outermost ones. Be careful not to get any medium on the plate between the wells.

27. The cells must be fed once a week during the next two lab periods by the addition of 0.1 ml of HAT medium to each well.

PART II: ISOLATION OF CELLS FROM ANIMAL TUMORS

OBJECTIVE:

To generate primary explants from B16 melanoma cells growing as tumors in C57Bl/6 mice.

PROCEDURE:

1. Use aseptic technique.

2. Surgical instruments will already be sterilized and should be kept in 70% alcohol once removed.

3. Select a mouse with a tumor that is at least 1 x 1 cm.

4. Sacrifice the mouse by cervical dislocation after anesthesia.

5. Soak the abdomen and side with 70% alcohol.

6. Carefully cut away skin exposing the tumor - do not touch intestines!

7. Remove the tumor using forceps and scissors and transfer to a petri dish containing 2 ml of normal growth medium for B16 melanoma cells.

8. Wash the tumor gently for 2 minutes.

9. To remove cell clumps and structural material pass the cells through a narrow mesh nylon screen by rubbing over the surface of the nylon that you hold over a petri dish. Keep the cells moist with medium. Wash the screen with 2 x 10 ml of medium.

10. Collect cells from the petri dish which have passed through the nylon screen and transfer to a 50 m

11. Spin down at 400 x g for 5 minutes.

12. Resuspend cells in 5 ml of ice-cold 0.87% ammonium chloride and incubate for 5 minutes.

13. Add 5 ml of growth medium then spin down cells and wash 2 x 5 ml with normal growth medium.

14. Remove an aliquot and determine cell counts and cell viability by trypan blue dye exclusion.

15. Set up a T-25 flasks with 5 ml of normal growth medium.

16. Observe cells 1 day after plating and look for attachment, cell morphology, the amount of pigmentation, and any other differences from normal B16 melanoma cells grown in culture. Record your observations for your lab report. These cells will not be used again so they should be discarded.

It is extremely important to be able to identify the cells that are grown in any culture, especially if cross-contamination or spontaneous transformation are possibilities. No single method is best for all cell types. General methods include the assay for specific cell functions (e.g. melanin production of melanocytic cells), cell-surface markers or antigens, specific growth requirements, and examination of cellular morphology (though the latter is less reliable and difficult to quantify). Analysis of chromosomal content (karyotyping) is one of the best methods for identification of cells. This method can be used to determine the chromosomal number and sometimes the specific banding patterns of individual chromosomes.

OBJECTIVE:

To learn the method of chromosomal analysis and to examine the chromosomes of murine cells.

CELLS:

1. Murine B16 melanoma cells

2. Mouse Skin fibroblasts (MSF-1)

3. Murine EG.7 thymoma Cells

PROCEDURE:

1. Flasks of all cells will be available.

2. Cells will already have been treated with colchicine.

3. Harvest cells by the normal trypsinization procedure.

4. Resuspend cells in 5.0 ml of hypotonic buffer (0.04M KCl, 0.025M sodium citrate) for 20 minutes at 37o.

5. Add 5.0 ml of ice-cold acetic methanol while constantly mixing then spin cells down at 200 x g for 3 minutes.

6. Buzz the cell pellet briefly on a vortex then slowly add 5.0 ml of acetic methanol while mixing and leave on ice for 10 minutes.

7. Spin down cells and resuspend cells in 0.2 ml of acetic methanol.

8. Remove 1 drop of the suspension and drop onto a slide, allowing it to run down the slide.

9. Dry the slide by placing it over a beaker of boiling water and examine it under the microscope. If cells are piled up and overlapping then the suspension must be diluted further until single cells are seen, then prepare a slide.

10. Immerse the slides in Giemsa stain for 2 minutes then run under water until the excess stain is removed (pink cloudy appearance).

11. Check staining under microscope and if satisfactory mount with glycerol.

12. Examine slides for chromosomal number and banding pattern if possible.

The ability to grow hybridoma cells in culture provides a tremendous opportunity to generate pure monoclonal antibodies as discussed in the laboratory on monoclonal antibodies. A major step in the procedure is the detection of antibodies needed to screen the hybridomas for antibody production. The ELISA methodology is used for this step. This technique is rapid, quantitative, and can be used to screen multiple (30 for example) microtiter plates. The method measures the binding of an antibody (found in the supernatant of the medium in the monoclonal method) to the antigen originally used to immunize the animals. The detection requires a second antibody that recognizes a mouse antibody. This secondary antibody is usually covalently linked to an enzyme (alkaline phosphatase is one that is used). Addition of the enzyme substrate allows us to detect the presence of the original mouse antibody that has been bound by the secondary antibody. The use of this enzyme-linked secondary antibody provides an extremely sensitive system.

ELISA methods are now routinely used in immunological assays testing for the presence of either antibodies or antigens. For example, cells grown in culture can be used as targets to screen for the production of monoclonal antibodies against those cells and this has been a very powerful tool of the monoclonal antibody technology. By analogy patients' sera may be tested against cultured tumor cells for the presence of antibodies against these cancer cells (or for antibodies against HIV with this as target). On the other hand (monoclonal) antibodies may be used to detect the presence of antigens on cells (either in culture or directly from tissue biopsies). There are several possible immunological methods, which can be utilized. In this laboratory the ELISA assay normally used to screen for monoclonal antibodies will be demonstrated.

METHODOLOGY:

The presence of antibodies reacting with soluble protein antigens will be analyzed. The antigen will be bound to all wells in 96 well microtiter plates by overnight incubation in coating buffer. The plate will be blocked with 5% milk in PBS for 1 hour. The plates can then be used to detect antibodies that may be present in the supernatant of your hybridoma wells.

OBJECTIVES:

1. To learn the basic methods of immunological assays using immunogens as antigen targets.

2. To determine whether antibodies against the immunogens were generated in the hybridomas made during the monoclonal antibody lab

ANTIBODIES:

Two positive control antibodies will be used to demonstrate that the assay works.

PROCEDURE:

1. Several important steps in the ELISA assay have already been performed because the assay would, otherwise, require too much time.

2. The plates will be blocked with 5% milk.

3. Remove the medium by inverting the plate over the sink and flicking your wrist, then blot the slide on a paper towel.

4. Wash the wells 2x with blocker (PBS containing 0.05% tween 20).

5. Add 100 ul of antibodies (from your hybridoma wells) diluted in blocker (5% milk) to designated wells.

6. Incubate the plates at 37o for 1 hour.

7. Wash the slides six times with tween-20 blocker.

8. Add 100 ul of secondary antibody (also diluted with blocker) and incubate at 37o for 1 hour.

9. Wash the slides six times with tween-20 blocker.

10. Add 100 ul of substrate solution to all wells and incubate at room temperature until color develops.

11. Measure the absorbance at 405 nm in the ELISA reader.

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