Making Monoclonal Antibodies
Monoclonal antibodies are great! Why? Specificity! Each one recognizes only one site of the antigen structure. Why else? Immortality! Mass cultures can be generated from a single clone.
Mice are small! They require less antigen than a rabbit to get an antibody reaction. The antigen travels through the circulation system. Blood is channeled through the spleen where B lymphocyte cells recognize the foreign bodies and produce antibodies.
Spleen cells, however, cannot survive alone in tissue culture media. Myeloma cells can! Why not fuse them?
We can select for fused cells by using a HAT (hypoxanthine, aminopterin, thymidine) supplemented media. The mutant myeloma cell line SP2/0 cannot survive in HAT media because it lacks HGPRT. The aminopterin in the HAT supplement blocks DNA synthesis. The enzyme HGPRT can overcome this block. Spleen cells do have HGPRT. Fuse the spleen and SP2/0 cells and you'll find survivors in HAT tissue culture media.
Remember, Sterile Technique!
Wash yo' hands.
If you pour (you really should pipet) DON'T SPILL! Contamination occurs because of spillage outside of bottles and tubes. Ethanol and flame when you're feeling paranoid.
For your own safety, wear gloves when dissecting.
Use disPo products to avoid problems with detergent residue on washed glassware.
BALB/c female mice for immunization
one mouse / feeder layer
disPo pipets - 10 ml and 25 ml
tissue culture flasks (5 / feeder layer)
24 well tissue culture plates (about 12 per fusion)
3 autoclaved sets of forceps and scissors
1 autoclaved screen for spleen cell dispersion
5cc syringe (plunger used to disperse cells on screen)
18g & 25g (1/2 inch) needles
100 mm tissue culture plates
15 & 50 ml sterile conical tubes
*see attached for details on ordering
Cutting out protein bands from a polyacrylamide gel.
Coomassie stain for an hour & destain (10% acetic acid, 45% methanol) as quickly as possible, cutting the bands as soon as they are discernible to limit the time that the gel is in acid. Rinse with PBSa & cut out the bands. Protein can be stored with a small amount of PBSa in the -20 freezer.
Make first injection of protein at 1:1 ratio with Freund's Complete or TiterMax.
1. Homogenize protein with a small amount of PBSa so protein doesn't get too thick. Keep protein cold while homogenizing.
2. If using Freund's, mix up dead bacteria in the bottle. Draw Freund's into a syringe and force it out into a small beaker containing your protein sample. Mix the Freund's and your protein until it takes on a white foamy consistency. You may have to change needles and syringes because the Freund's reacts with the plastic.
e.g. To inject 6 mice with emlc and rmlc each, I ran one 12% gel, cut out the bands, homogenized the protein and combined each protein sample with 450ul of Freund's. This gave me a little over 600ul of each sample to inject.
3. To inject. Refer to the animal care manual for further instructions.
· Inject 100ul per mouse.
·Your sample must be concentrated but not so thick that you cannot get it through a 1/2 inch, 25g needle.
·Sterility is important for the health of the mouse!
Two weeks later, make second injection of protein with Freund's Incomplete Adjuvant or TiterMax.
Two weeks later, do a test bleed. If negative, do another boost with Freund's Incomplete Adjuvant or TiterMax.
Five days before the fusion, do a final boost of protein without Freund's.
Mark mice's tails.
Apply grease to tail (an area of 3/4 inch, from base toward tip).
Use extremely sharp blades to make slice (EM blades). Be careful not to cut through the tail but make cut large enough to draw about 2.5 heprinized capillary tubes.
Apply pressure to tail wound to stop the bleeding.
Spin down the blood at max speed for 5 minutes in the microfuge.
Draw off and store sera at -20o until you can test on a blot at 1:100 or at 1:50.
Preparation for Fusion Day
Thaw out a plate of SP2/0, split one to 3.(must split during weekend). Need 10-12 100mm plates in log phase for fusion.
Thaw tube quickly in 37o bath. Put into falcoln tube with 10ml of warm media. Let sit for 5 min. Spin down at low speed. Resuspend in 10 ml and spin. Resuspend in 30ml and put into 3 plates.
Run gel for final boost.
Do final boost of protein without Freund's or Titermax.
Prepare Feeder Layer
FEEDER LAYER DAY
animal board- ethanoled and then sterilized under UV light
2 x 10 ml petri dishes
autoclaved forceps, scissors, & screen
5 ml syringe, 10 ml syringe, 18g needle
beaker with 95% ethanol
disPo 10 ml pipets
Warm the following in the 37o water bath:
·50 ml aliquots of DMEM- without additives
·500 ml bottle of DMEM two which the following has been added:
5 ml Sodium pyruvate
5 ml L-glutamine
HAT(with syringe, pump 5ml DMEM into HAT bottle, add to DMEM)
5 ml anti-biotic/anti-mitotic(Don't warm past room temp)
50 ml Hyclone Fetal Calf Serum
Put into 5 x 50 ml conical tubes (to later be transferred to 5 flasks for feeder layer).
· Set up the sterile dissecting utensils.
· Get the normal un-immunized mouse.
· Place screen in plate with 10 ml of DMEM-.
WORK QUICKLY! (The slower you are, the more difficult it is to work with the animal. Also, you don't want the media to cool.)
· Sacrifice the mouse. Ethanol until saturated. Pin the animal in place.
· Use one set of utensils to cut through the skin, pin back the flap. ETHANOL!
· Use second set of utensils to cut through the peritoneum.
· Use third set to pull out the spleen, cutting away the connective tissue.
Changing utensils is very important to avoid contamination.
·Put spleen in the dish with 10 ml of DMEM-. Quickly and carefully disperse spleen cells with plunger of a 5 ml syringe against the screen.
·Leaving connective tissue on the grid, pipet up and down and put cells into a 15ml conical tube. Let large chunks settle to the bottom of the tube for 2-3 min.
·Transfer supernatant to another tube bringing the volume up to 15 ml with more DMEM- to wash. Spin down for 10 min at 170g on the table top centrifuge.
·Carefully, suck off supernatant with sterile pasteur pipet. Wash spleen cells again with 10ml DMEM-.
·Suck off super. Resusp. in 5ml DMEM-, incubate for 10 min. in 37o bath. Pour 5x 50ml conicals of DMEM+ into tc flasks. Put 1ml of spleen cells in each flask.
·Incubate o/n in 6% CO2 37o chamber. Lay flasks on sides with caps loosened.
I. Collect immunized spleen cells.
II. Collect SP2/0 cells.
III. Combine with feeder layer.
· All of the same supplies used for the feeder layer on previous day.
· Transfer 50% PEG into a conical tube.
· Warm the PEG, wash media, and the remaining DMEM+ in 37o water bath.
Filter sterilize leftover DMEM+ media from yesterday followed by feeder layer. The feeder layer is more likely to clog the filter.(Make sure you have media in the container before turning on the vacuum otherwise you could break the filter membrane).
Prepare the Immunized Spleen Cells
Follow protocol used to dissect the mouse for the feeder layer.
It is important to minimize the time that the spleen cells are either not in the animal or not in the incubator. For this reason and depending on how quickly you are able to dissect the mouse, you may want to collect the SP2/0 cells first.
Collect the SP2/0 Cells
Triterate cells from 10 plates and put into 2 x 50 ml conical tubes.
Spin down the cells. Wash cells 2x. Pellet for 10 min. at 170g.
|pellet spleen cells||WHILE||triterating the SP2/0 cells|
|resusp spleen cells, wash, pellet||WHILE||pellet SP2/0 cells|
|resusp spleen cells, wash, pellet||WHILE||resusp one tube in 5ml, transfer to |
other tube, wash, pellet
|resusp spleen cells in 10 ml, count,||WHILE||resusp SP2/0, wash, pellet |
let sit for 10 min in 37o
resusp in 20-30 ml, count
|1 x 108 spleen cells||ADD TO||5 x 107 SP2/0 cell|
· Pellet cells together at higher speed (between 170 - 200g).
· Suck off media
· Add 1.5 ml PEG over 1 min. under agitation-- THIS IS THE FUSION!!!
Stir gently with tip of pipet to mix in PEG
· Wait 90 seconds, stirring gently.
· Over the next min, add 1ml of warmed DMEM- while stirring.
· Repeat previous step.
· Add 8 ml DMEM- over 5 min.
· Wait 10 min. Then, disrupt cells by pipetting gently 5 times.
· Dilute into sterile filterized DMEM+ with Feeder Layer.
· Aliquot 2ml/ well in 24 well plates (~ 10-12 plates)
· Incubate in 6% CO2, 37o chamber for one week.
· Feed and incubate for ~one week.
one week after first media feeding:
Test wells where media is turning yellow.
Draw off 500ul & test on blot strips, incubate overnight. (Don't forget positive controls including mouse antibody - use mouse polyclonal sera and a marker.)
Grow up positive wells in larger plates.
Retest the well and its corresponding plate.
Both should be positive. Continue to split positive plates and freeze down (in 10% DMSO) as back ups. Triterate and dilute cells from the positive well into a 96 well plate at a dilution of 1 cell to ~3 wells. We're going for the single clone!
Transfer cells in the 96 well plates to 24 well plates and Retest.
Pass through dilution again until all wells are testing positive and you are convinced that you have a monoclonal cell line.
Eventually, try and wean the cells from 10% FCS to 5% FCS, and from HAT to HT to unsupplemented media.
Affinity Purification of IgG Using G-Beads (Pharmacia)
0.1M NaPO4, pH 7.0
monobasic - 5.52g in 200 mL for 0.2M monobasic
dibasic - 8.52g in 300 mL for 0.2M dibasic
start with the dibasic solution and titer with the monobasic until pH 7.0 is reached
dilute 1:1 with water for use
1.0M acetic acid - 0.1M glycine buffer, pH 3.0
30.12 mL acetic acid
3.75 g glycine
bring to 500 mL with water
1. Equilibrate column with several volumes of pH 7.0 buffer.
2. Put sample over column - 10 mL rabbit serum for IgG isolation or entire volume of papain digestion - take gel samples. (Flow rate at 1mL/4 min)
3. Save pass through - in papain digest the pass through contains the Fab - take gel samples.
4. Wash to baseline with pH 7.0 buffer (1-2 hours).
5. Elute with pH 3.0 buffer - takes 15-20 minutes to see something.
6. Collect 0.5 mL fractions and neutralize solution with 1.0M Tris to pH 7.4 (use indicator strips)- take gel samples. Spec fractions to find peak.
7. Wash with pH 3.0 buffer for 15 minutes and store in 20% ethanol.
8. Dialyze IgG or Fab against 4L PBS overnight at 4° C then at room temperature with 2 changes over 1-2 days.
9. Determine concentration using spec at absorbance of 280nm with A280 of 1.35 for IgG or 1.5 for Fab.
10. Store at -80° C in aliquots.
1. Run SDS-PAGE curtain. (Use as much protein as possible without having so much that you risk running bands together and contaminating the band of interest).
2. Place unstained gel in 0.5 M KCl for 10-15 seconds -- until bands begin to appear. The bands will appear clear, while the rest of the gel will turn white as the SDS precipitates. Remove the gel onto a glass plate and cut out band of interest as rapidly as possible. If you wait too long, the bands will turn white as well.
3. Place gel strip(s) into 50 mM NaPO4, pH 6.5 with 0.1% SDS and rinse 3 times, with gentle shaking, for 30 min. per rinse.
4. At third gel wash, begin activation of the APT paper. Do these steps in the cold room on an ice bath. First, place an appropriately-sized piece (slightly larger than gel) of APT paper in cold 1.2N HCl (100 ml) to which has been added 3 ml of sodium nitrite (NaNO2). The NaNO2 is made in water at 10 mg/ml immediately before use. Incubate paper on ice with occassional shaking for 15-30 min. The paper should turn bright yellow. Activation is followed by two or more rapid ice water washes and two rapid washes with 50 mM NaPO4 pH 6.5, taking 5 minutes for the whole process. (When I originally did this, I used a protocol which called for 5 X 5 minute washes with ice cold water and a subsequent 10 min wash in cold 50 mM NaPO4, pH 6.5. However, Bio-Rad emphasizes the necessity of short washes. Both seemed to work for me.)
5. Immediately after activation, set up the transfer in the cold using 50 mM NaPO4, pH 6.5 as transfer buffer. Transfer for 4 hours at 0.6 amps (Hoefer TE-52). The paper should have turned an orangey (peach) color. Ususally where the gel was, the color is lighter.
6. Block the DPT paper in the following buffer for 2 or more hours:
0.25% gelatin (0.5 g) dissolve first by heating
20 ml ethanolamine
20 ml 1 M Tris-HCl pH 9.0
to 200 ml with dH
7. Rinse extensively in "Buffer 1" following the blocking step. Do at least 3 20-30 minute washes.
50 mM Tris-HCl, pH 7.5
5 mM EDTA
150 mM NaCl
8. To check transfer efficiency, cut a strip off one side of DPT paper to which protein has been transferred and incubate with your (appropriately diluted) antibody overnight. (Dilute the antibody in Buffer 1 containing 0.25% gelatin, or 1% BSA [Sigma Fraction V]. I use BSA.) Carry through with normal protocol for immunobloting, i.e. rinse (3 changes of buffer 1 or PBS) 30 min, incubate 2-4 hours in peroxidase conjugated secondary antibody, rinse, develop in 0.05% 4 chloro-1-napthol (made up as a 0.3% solution in methanol), 0.01% H2O2 in PBS. The reaction seen on the DPT paper is usually not as distinct as that seen on nitrocellulose.
9. If you are waiting to check efficiency of transfer before using the DPT paper, just store paper in Buffer 1 at 4o C until use.
10. To use antigen-bound DPT paper:
a. cut paper into small squares (on a glass plate) with a clean razor blade.
b. place squares in 10 ml syringe with large gauge needle
c. incubate with 5 ml of an appropriate dilution of antiserum (I use a dilution 2 times more concentrated than that used for immunoblotting). Dilution is made in Buffer 1 with BSA or gelatin. Incubate overnight with vigorous shaking. I do this at R.T. without any problems
d. the next morning, squirt out diluted antiserum (from which epitope specific antibodies have presumably been removed) and draw about 10 ml of Buffer 1 into the syringe. Shake rapidly and repeat for a total of 3 rinses (30 minutes each)
e. to elute specific antibodies, use 4 ml of 5 M NaI made up fresh in ice H2O. Draw up into syringe and place on ice; agitate by rotation of the syringe on ice for 8 minutes
f. force the NaI solution (now containing eluted antibodies) into a tube (50 ml Falcon type) containing 600 ll of Buffer 1/1% BSA. Force Buffer 1 through DPT paper repeatedly, pooling all of these washes in the same 50 ml tube until a total of 40 to 50 mls is reached. (You may have to dilute even further if the intended use of the antibody is for immunofluorescence. Concentrations of salt that are too high interfere with fluorescence visualization).
g. To concentrate antibody:
Concentrate using a Millipore immersible CX-10 filter. Prepare the filter by rinsing with dH
Alternative #2. (faster)
a. prepare two amicon centricon filters (30,000 mw cutoff) by adding 300
b. place one half the 5M NaI/antibody solution in the upper reservoir of each of two centricon filters. Using a pasteur pipette, stir to mix the two solutions.
c. spin at 5000 x g for 30 minutes. (Use parafilm to make the filters to fit into rotor adaptors.
d. after the spin, discard the liquid which has passed through the filter (i.e. the liquid in the lower reservoir).
e. bring liquid remaining in the upper reservoir (100-500ml) to a volume of 2 mls with buffer 1.
f. spin again at 5000 x g for 30 minutes.
h. discard liquid which has passed through the filter.
i. add 250ml of buffer 1 to each upper reservoir and cap with storage reservoir.
j. invert tubes and spin in table to centrifuge for 2-3 minutes to transfer the entire sample into the storage reservoir.
NB. filters can be reused for repeated purification of the same antibody. Store at room temperature with 1 ml of buffer 1 in the upper reservoir.
11. Store DPT squares in the syringe in Buffer 1 at 4oC. I've never used azide but have used squares repeatedly (up to 10-12 times) over a period of 6 months without any problems.
1. Harvest 5 x 106 - 4 x 107 cells/sample depending on how abundant your protein is. Use 5 x 106 cells/sample to IP myosin, 4 x 107 cells/sample to IP dynein, for example. Wash cells a couple of times in DB, and resuspend in 500 m l DB/sample. Place on ice.
2. Add an equal volume 2X IP buffer (made fresh) and mix on ice:
2X buffer for dynein IP:
100 mM Pipes, pH 7.5
10 mM EDTA
50 mM NaPPi
200 mM NaF
5 mM DTT
2 mM PMSF
100 m g/ml leupeptin
100 m g/ml pepstatin
2 mM ATP
2X buffer for myosin IP:
40 mM Tris, pH 7.5
2 mM DTT
10 mM EDTA
2 mM PMSF
200 m M TPCK
200 m M TLCK
20 mM NaHSO3
50 mM NaPPi
200 mM NaF
2 mM ATP
200 mM KPO4
3. Add 100 m g/ml RNAse A.
4. Centrifuge lysate 18K, 30 min, 2 degrees, in SS34 rotor
5. Collect cleared lysate. Use 1 ml lysate per IP sample. Add primary antibody:
For NW127 dynein HC Ab, add 50 ul serum/sample
For 142, 143, 144 dynein IC Abs, add 50 ul serum/sample
For 9E10 anti-myctag Ab, add 200 ul supernatant/sample
For 396 myosin Ab, preincubate 25-30 m l GammaBind or protein A-sepharose beads with 1 ml of 396 supernatant for 4 hours to overnight. Wash beads as per steps 10-11 below and resuspend beads directly in 1 ml cleared lysate. Skip steps 6-7 and go directly to step 8.
6. Rock in cold room for 2 hours to overnight.
7. Add 25-30 ul protein A-sepharose slurry for rabbit and rat primary antibodies; add 25-30 ul GammaBind slurry for mouse primary antibodies.
8. Rock in cold room 1-4 hours.
9. Spin down beads at 5K for 2 minutes in microfuge.
10. Wash beads twice with 1 ml each time of 1X IP buffer with 1 mg/ml BSA (to reduce background, adjust pH of IP buffer to 7.0 for washes). Spin down as above.
11. Wash beads twice with 1 ml each time of 1X IP buffer, pH 7.0 without BSA. Spin down as above.
12. If desired, do a final wash in 1X IP buffer with 0.5M NaCl. This will further reduce your background. HOWEVER, it will strip the dynein IC off the HC when doing IPs with NW127 Ab!
13. Remove all traces of wash buffer, and resuspend beads in 20-30 m l 2X SDS sample buffer. Boil 5 minutes. Spin down beads, load super onto an SDS gel. IgG heavy chain runs at 50 kd; IgG light chain runs at 22 kd.
notes: If you have a lot of background, it may help to preincubate the cleared lysate with beads (without antibody!) for a couple of hours to adsorb out any non-specific binding. Spin down the beads, recover the lysate, and add primary antibody. Proceed with step 5.
Specific Details for Myosin Ips
1. Cells were harvested and washed in Ca2+-free MES starvation buffer (20 mM MES, 2 mM MgSO4).
2. Ice-cold 2 x lysis buffer was then added to the cells suspension. (40 mM Tris-Cl pH7.5, 0.2% NP40, 2mM DTT , 10 mM EDTA, 2 mM PMSF, 2 mM TAME, 200 m M TPCK, 200 m M TLCK, 20 mM NaHSO3, 100 m g/ml RNAse A, 50 mM Sodium pyrophosphate, 200 mM NaF, 2 mM ATP, and 200 mM Phosphate pH7.5)
3. Vortex and then centrifuged for 30 min at the top speed of the microfuge.
4. The supernatant was transferred to a tube containing mAb and 20 m l 1:1 protein A sepharose slurry, and the tube was rotated end-over-end overnight at 4 oC.
5. The immunoprecipitate was then washed at least 6 times with MES-Salt buffer(20 ml MES pH 6.8, 20 mM NaCl, 1 mM EDTA) for 5 min. each.
6. The beads were pelleted for 2 min. at a microfuge, resuspended into 2 x SDS sample buffer, boiled for 5 min. at 90 oC sand bath.
7. The slurry was pelleted again, keep and supernatant.
8. Run SDS-PAGE.
IgG heavy chain will run at 50kd
IgG light chain will run at 22kd
Whole IgG runs at 150kd
Need to worry about detergents used to lyse cells inhibiting Ab binding.
Need to worry about immunocomplex not being soluble.
Need to work in zone of equivalence (Ag:Ab ratio).
Instead of 2° GAMIgG can use whole Staph. aureus, Protein A, Protein G which binds to IgG1 better than Protein A, Prot.A/Prot.G mix (Pierce Immunochemicals), or antimouse Affigel.
Can release antigen from immunocomplex by 0.1M Glycine pH 2.5 acid shock, or high salt (3M NaSCN) shock, then passage through Costar Spinex 0.22m m filter unit into Eppendorf.
Can increase titer of 1° mAb by passing 9E10 media onto Sepharose CNBr beads, this binds 5 or more Fc portions of the IgG effectively making it an IgM.
Can increase titer of 1° mAb by adding equal volume of saturated NH4SO4 dropwise to spinning 9E10 media at roomtemp, pelleting the precipitate at 2500g for 5min, resuspending the pink (RPMI) pellet in dH20, repeating the precipitation.
To make saturated NH4SO4: boil water, add NH4SO4 until crystals fall out of solution, cool to RT, pH to 7.0 with NH4Cl, filter through 3MM paper.
Can grow 9E10 cells in 2% FCS to lower contaminating Ig.
This protocol takes advantage of the fact that antibodies transfer rapidly between antigen samples attached to separate solid supports. It allows one to affinity purify small amounts of antibody using less than a picamole of antigen. This is great for doing immunofluorescence with dirty antibodies that are in short supply or whose antigens are in short supply, or for quickly screening expression clones for epitope selection.
1. Immobilize your antigen on a 1 cm2 chip of nitrocellulose. For example, spot blot 1-10 ng of native Dicty dynein in 10 m l P100 buffer onto a nitrocellulose chip. This is your source blot. Make one source blot for every coverslip you wish to stain.
2. Let nitrocellulose dry. Mark the protein side with a pencil. Block in 5% milk/PBS for 30 minutes. Source blots can be stored in blocking solution in the fridge if NaN3 is added to 0.02%.
3. Incubate antigen source blot with antibody in PBS/0.1%BSA for 4 hours to overnight. Handle blots with blunt forceps. Use a dilution of antibody that gives good results with a Western blot. The wells in a 24-well plate are good for this if you have several samples.
4. Wash source blots 3 times 5 minutes with PBS/0.5% Tween 20.
5. Prepare coverslip sample for indirect immunofluorescence (see relevant section in labman). Place 50 m l PBS on fixed, blocked coverslip. Place washed source blot, antigen/Ab complex down, on top of coverslip. Incubate at 37 degrees, 1 hour, in humid chamber. The antibody on the source blot will rapidly equilibrate with the antigens on the fixed cells on the coverslip. Proceed with washing and secondary antibody as per immunofluorescence protocol.
6. Source blots can be re-used if washed extensively and stored in PBS/0.1% BSA/0.02% NaN3. Re-incubate with antibody each time.
For epitope selection of expression clones
1. Grow up dense IPTG-induced (clonal!) phage plates and lift expressed fusion proteins onto nitrocellulose filters as usual.
2. Block phage filters in 5% milk/PBS for 30 min. Incubate with the antibody for 1-4 hours at the same dilution used to screen the phage library. Wash filters in PBS. These are your source filters.
3. Run a curtain SDS gel of a sample of purified or enriched antigen. Blot the gel onto nitrocellulose and mark the protein side with a pencil line across the entire width of the blot. Block in 5% milk/PBS for 30 minutes and cut into strips.
4. Incubate each source filter to be tested with a strip of curtain gel. Place the source filter and the blot strip with protein sides facing each other in a seal-a-meal bag. Add just enough PBS/0.1%BSA to wet both blots. Seal the bag. DO NOT ADD ANTIBODY AT THIS POINT!
5. Incubate blots together for 1-4 hours. Then remove blot strips and wash extensively in PBS. Do not mix strips in washing solution or secondary antibody solution, as antibody may transfer between strips. Incubate the strips in the appropriate secondary antibody for 1 hour. Wash and develop strips.
6. If a phage clone encodes a truly positive cDNA, the corresponding strip will be positive as well. Negative strips indicate false positive phage clones.
(From Hammarback and Vallee ,1990, JBC 265:12763)