Shreffler:Milk Program

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Revision as of 07:41, 31 July 2009 by Alyssa Chase (talk | contribs) (Materials)


Milk allergy is the most common cause of food allergy in infants and young children, affecting 2.5% of infants in industrialized countries. Previous studies suggest that the majority will tolerate heat-denatured products without harmful effects. Our preliminary findings that ingestion of extensively baked-milk products may enhance tolerance induction. This study is designed to determine whether frequent dose escalation of milk protein in allergic patients would increase the tolerance of their immune system.

Through the baked milk 2 study, we plan to study the immunologic mechanism associated with oral tolerance induction. We propose to investigate the effect of introducing baked-milk products by comparing the rate of tolerating less heated milk in children subjected to more frequent dose-escalation (every 6 months) and in children subjected to less frequent maintenance increases (every 12 months).There will be a comparison of the tolerance of an individual patient longitudinally over a period of time. Also, the changes in T regulatory cells would be evaluated in five groups, who are given different doses of baked milk proteins.

Day 0 Procedure: Isolation of PBMC's, Basophil Activation Assay, CFSE Labeling, T Regulatory Cell Assay, & Antigen Activation Assay


Isolation of PBMC's (General protocol described ____)

  • 18 mL sample of whole heparinized blood/patient
  • Ficoll Paque Plus [endotoxin tested], room temperature
  • Sterile phosphate buffered saline (PBS), room temperature
  • 0.2% and 0.4% Trypan Solution
  • Sterile conical tubes (15 mL, 50 mL)
  • Sterile, graduated transfer pipettes
  • Hemocytometer or disposable slides for the automated counter

Basophil Activation Assay (General protocol described: ___)

  • 3 mL sample of whole herparinized blood/patient
  • RPMI medium (store at 4°C in dark)
  • 1x FACS Lysing Solution (made from 10X stock w/dH2O, store at 4°C, expires in 1 week)
  • Staining Buffer (PBS + 2 mM EDTA + 0.5% BSA) (sterile filter, store at 4°C; aliquot in hood; expires in 2 months)
  • Monocolonal Antibody cocktail (CD63-FITC, CD123 PE-Cy5, HLA-DR-PE Cy7, CD41a-APC, CD3-APC, CD14, CD19-APC) (store at 4°C in the dark).
  • Polypropylene tubes
  • Pre-made 30 μL aliquots (stored at -30°C):
 a.  Basophil medium w/ 20 μM of fMLP
 b.  Basophil medium w/ 200 μg/mL milk protein
 c.  Basophil medium w/ 20 μg/mL anti-IgE
 d.  Basophil medium w/ 20X PMA (5 μg/mL )
 e.  Basophil medium w/ 20X Calcium Ionophore (20 μg/mL) 

  • 15 mL polypropylene tubes
  • PBS
  • 5,6-CFDA/SE Invitrogen C1157 dissolved in DMSO at 5 mM and stored at -20°C.


Basophil Assay and Isolation of Mononuclear Cells (done simultaneously)

  1. Label 4 (50mL) tubes as PBS: Blood, AIM-V, PBMC, Ficoll and 2(15mL) tubes as Basophil blood and RPMI
  2. Bring 15/10mL of Ficoll(sterile) to room temperature and put 5mL RPMI in water bath.
  3. Collect pre-made stimulants from the freezer (Rm 40 in clear boxes with rubber bands). LABEL each one
  4. Collect 6 epindorph tubes. Label 1 of these tubes "basophil cocktail." Make a cocktail as follows: Combine 180 μL RPMI + 3.6 μL IL-3. MIX WELL.
  5. Label the 5 remaining tubes "Basophil medium." Add 30 uL of the cocktail into these tubes.
  6. Add 270 uL of the warmed RPMI to the five basophil tubes(made above), fMLP, milk extract, and anti-IgE.
  7. Add 135 uL of RPMI to PMA and CaI and then combine them.
  8. Prepare a 10-fold dilution of milk stimulants:
  • Transfer 270 uL of basophil medium from the aliquots prepared in step 2 to each of the milk 2-5 epindorph tubes.
  • Transfer 30 uL from tube "milk 1" to "milk 2". VORTEX.
  • Take 30 uL from "milk 2" and add to "milk 3" etc. Vortex after each step.

9. Label 5 mL polypropylene tubes A-K. Remember to include subject number on tube A. list of stimulants

10. Transfer 250 μL of warm RPMI to tubes A & B.

11. Transfer 250 μL of each stimulant to the appropriate polypropolene tube (C-K) and keep them in incubator until ready to start.

12. Get the blood from the clinic, open the two green top tubes under the hood.

13. (keep sterile) Set aside 3mL of blood into pre-labeled tube for basophil assay.

14. Transfer remaining blood in pre-labeled 50mL tube and dilute 1:1 with PBS

15. Overlay with 30 mL of diluted blood on the 15mL ficoll.

16. Centrifuge at 500 g for 30 minutes at 23°C with acceleration slow and brake off. now back to basophil protocol

17. Take out the labeled tubes from incubator

18. Transfer 250 μL of participant blood to each tube A-K.

19. Incubate tubes for 30 minutes at 37°C (in incubator).

20. Prepare the cocktail. Label a 5 mL polypropylene tube "staining cocktail" Add 700 μL of staining buffer.

   a. 35 uL of HLA-DR-PE-Cy7 mAb
   b. 70 uL of CD63- FITC
   c. 70 uL of CD203c-PE
   d. 70 uL of CD123 PE-cy5
   e. 70 uL of CD41a
   f.  70 uL of CD3
   g. 70 uL of CD14
   h. 70 uL of CD19-APC

21. Remove tubes from incubator and add 50 μL cold PBS w/20 mM EDTA to each tube to stop degranulation.

22. Stain cells by adding 110 μL of the prepared Ab Cocktail to tubes B-K. Do NOT add to Tube A.

23. Put it in the refrigerator(4°C) for 30 minutes.

  • Return to isolation of PBMC's. Remember to keep conditions STERILE!

24. Remove ficoll tubes from centrifuge. Using a sterile transfer pipette , collect PBMCs (cloudy gray layer) into a new 50 mL conical tube.

25. Under the hood, add sterile PBS to double the volume and invert the tube to mix. Centrifuge at 500 g for 20 minutes at room temperature (max acceleration and deceleration).

26. Aspirate and discard the supernatant. Resuspend the pellet by tapping the tube until no clumps are visible, then adding 1 mL of PBS.

27. Set aside a 10 μL aliquot of cells for counting: 10 μL of cells into 90 μL of PBS in a STERILE eppendorf tube.

28. Add PBS to cells to make a total volume of 20 mL.

29. Centrifuge at 300 g for 15 minutes at room temperature (maximum acceleration and deceleration).

30. While the cells are in the centrifuge, you must determine the volume to use for resuspending the PBMCs after this wash. This in turn requires knowledge of the total number of cells in the sample:

a. Combine the 100 μL aliquot of cells in PBS set aside in Step 28 with 100 μL of 0.2% Trypan solution (if using the automated counter) or 0.4% Trypan soplution (if manual counting).

b. Mix well with a pipette.

c. Place 20 μL of the stained cells onto a disposable slide.

e. To take the cell count, in the computer program, write dilution as a whole number (20x). [1st diluted 10 μL of cells in 100 μL total volume PBS + Cells, then added an additional 100 μL of Trypan solution

           = 10μL/200μL = 1/20 = 20 fold dilution).

f. Choose cell type from drop down menu ("Human")

g. Pick "display"

h. Focus image so the cells appear yellowish

i. Hit "Count."

31. After centriguation is completed, aspirate and discard the supernatant. Resuspend the cell pellet by tapping the tube until no clumps are visible. Suspend PBMCs at 10 x 10^6 cells/mL in PBS. To calculate this: Take number counted in step 31i divided by 10 x 10^6 cells/mL. This gives you the the total volume of PBS you must add to the cells.

32. Divide Cells as follows: a. 10 x 10^6 cells for measurement of Th1/Th2 gene transcription in allergen-activated cells by RT-PCR (after 48 hr culture). b. 10 x 10^6 cells for measurement of frequency of PBMC-derived, milk-specific CD4+, CD25+, FoxP3+ T cells (after 7 day culture). c. If indicated, remaining cells will undergo CD25 Depletion Experiments.

  • Note, if you collect less than 2 mL of PBMC's in step 32, just divide the volume equally to the 48 hour and 7 day test tubes in step 33.

Now prepare a T-Regulatory Cell Assay for 48/ 7 DAY CULTURE PREPARED ABOVE.

33. In a 15 mL conical tube, make a PBS: CFSE solution: a. 1 mL PBS b. 2 μL of 5 uM CFSE

34. Add the PBS: CFSE solution just prepared to the 7 DAY CULTURE PBMC's.

35. Place in a 37°C water bath for 10 minutes.

Proliferation Assay

36. STERILE CONDITIONS. Label two 24 well plates/their lids with specimen ID and date. Label each well with the appropriate condition, ordered by priority (for cases where there are insufficient cells to test all stimulants). a. Allergens- Caseins (C+): 50 μg/mL purified casein proteins (alpha, beta, kappa) in AIM-V medium b. Negative Control- AIM-V medium (A+): Medium alone c. Positive Control- Beads (B+): 1 μg/mL anti-CD3, anti- CD28 beads in AIM-V d. Egg white (E+): 20 μg/mL egg white in AIM-V

37. Label one plate for 48 hour culture and a second plate for a 7 day culture.

38. Add 10 mL of AIM-V to both 7 and 48 hour culture tubes and spin at 300 g for 10 minutes at room temperature.

39. STERILE CONDITIONS: Aspirate the cells.

40. Under the hood, resuspend cells in 2.5 mL of AIM-V medium to obtain a concentration of 4 x 10^6 cells/mL (For plating, each well should contain at 2-2.5 x 10^6 cells and a total volume of 1 mL).

41. Prepare solutions for each stimulant condition in sterile, 5 mL polypropyene tubes. Label each tube with a notation for 48 hour culture and A+, B+, C+, E+. Also, prepare tubes for the 7 day culture samples and also label A+, B+....

42. 48 Hours Antigen Stimulation Preparation a. Place 500 μL of AIM-V in each of the test tubes. b. Add 500 μL of the 48 hr-labeled cells in AIM-V to each tube and mix by pipetting up and down. c. Then add to the respective tubes:

A+: Medium Alone. Do not add any other substances.

B+: Add 5 μL of CD3, CD28, being sure vortex first to resuspend the beads in their container.

C+: Add 2.5 μL of EACH casein (alpha, beta, kappa).

    • Be sure to mix by pippetting up and down.**

43. 7 Day Antigen Stimulation Preparation: Unlike 48 hour culture, this contains CFSE labeled cells and IL-2.

a. Create a cocktail. Place 2 mL of AIM-V + 4 uL of IL-2 in an appropriately labeled test tube. Vortex gently. b. Add 500 μL of the cocktail to the labeled A+, B+, etc. test tubes. c. To each tube, add 500 μL of the 7 day cells set aside in step 33. Mix. d. Then add to the respective tubes:

A+: AIM-V medium + IL-2 alone. Do not add any other substances.

B+: Add 5 μL of CD3, CD28 expander beads. Be sure to vortex the beads in their container. Then suspend the beads in the AIM-V/IL-2 solution by pipetting up and down.

C+: Add 2.5 μL of each casein (alpha, beta, kappa). Pipette up and down.

E+: Add 10 μL of egg white. Pipette up and down.

44. Plate the stimulants for both the 48 hr and 7 day samples into the wells labeled in step 37. Then place the tissue culture plate in the incubator.

Procedure for Splitting Cells with 7 Day Incubation Period

  If you receive a sample on Monday or Tuesday, splitting should be completed on Friday.  For Wednesday samples, split on Monday.

1. Obtain the appropriate plate. Set pipette to 250 μL. Mix by pipeting up and down in each well.

2. Take 250 μL of culture from the well labeled A+ and add it to the three wells vertical to it. Repeat with conditions B+ through E+.

3. Add 750 μL of AIM-V at ROOM TEMPERATURE to each well.

48 Hours Protocol

1. Obtain the plate labeled "48 hours" that was placed in the incubator during the Day 0 procedure.

2. Label four 5 mL polysterene tubes (A+, B+, etc.). Remember to have the ID of the patient on the first of the tubes. Collect the specimen from the incubator dated two days before the date the 48 hr procedure is performed (obviously).

3. Label 12 CLUSTER TUBES as follows:

a. Specimen ID

b. Date of original culture

c. 4 tubes-- Supernatants, 48 hr.

d. 4 tubes-- Cx Cells, 48 hr.

e. Remaining tubes- JUST label ID, flush, A+, B+, etc.

4. Resuspend cells by pipetting up and down (getting the "four corners" of the well), and then placing the fluid in their respective tubes. Be sure to transfer the total volume/well.

5. Centrifuge tubes at 300 g for 5 minutes at room temperature.

Go to the "Collection of Supernatants Protocol."

6. Obtain a cluster rack for the storage of supertanants.

7. For each stimulant, using a 1000 uL pipette tip, transfer 800 uL of supernatant from the culture tube into each corresponding cluster tube int he cluster rack. Be careful not to disturb the cell pellets.

8. Cap the cluster tubes and store in the -80 C freezer. Keys for the freezer are on a blue chain by the lab bench. Our box is in the top fridge, bottom right, and it is labeled "milk project."

Return to 48hr procedure.

9. Shake tube gently to dissolve pellet.

10. Obtain 30 mL of Running Buffer (In the fridge Rm. 46), this is to be used throughout the experiment.

11. Add 1 mL of Running Buffer (a type of Cell Separation Buffer) to the dissolved pellet tubes.

12. Centrifuge at 300g for 5 minutes. Aspirate.

13. Resuspend cells in 80μL of cell separation buffer.

14. Obtain anti-CD25 Microbeads from the small yellow and blue box from the fridge in Rm 46.

15. IN THE HOOD add 10 μL of anti-CD25 Microbeads to each sample.

16. Mix well and incubate for 15 minutes in the refrigerator.

17. Cool the Centrifuge to 4C (but do not spin anything).

18. Wash cells by adding 1.5 mL of cell separation buffer and centrifuge at 300 g for 10 minutes.

19. Aspirate supernatant completely.

20. Resuspend cells in 500 μL of cell separation buffer. (PAY ATTENTION TO UNITS!!)

21. Obtain the green and black magnet (OctoMACS magnet) from the large cabinet under the bench.

22. Obtain Macs separation columns (Also called MS columns) from the supply station in the back of Rm 46. Be sure to keep columns on paper towels, not on the bench.

23. Place one MS column for each of the give cell cultures onto the OctoMACS magnet.

24. Underneath the columns, place 5mL polypropylene tubes.

25. Flush each column with 500 μL of cell separation buffer.

26. Apply cell suspensions to the corresponding columns.

27. Add 1 mL of cell suspension buffer to the original tubes to wash.

28. Once the column reservoir is empty, remove 500 μL of buffer from the original tubs and apply to the columns 2 times. (500 μL each time)

29. Remove columns from the magnet and place tip into an appropriately labeled eppendorf tube.

30. Pipette 1 mL of buffer into the column.

31. IMMEDIATELY flush out the fraction with magnetically-labeled CD25+cells by firmly applying the plunger supplied with the column. To do this, hold the eppendorf tube and bottom of column in left hand. Make sure injector is far from the bottom of the tube to avoid flood. Then tighten on the plunger. When you push, make sure to give the fluid room to flow.

32. Using a MICROCENTRIFUGE collect the pellets. Turn the eppendorf tubes so that the opening is facing the middle of the centrifuge. Be sure to balance.

33. Microcentrifuge at 400 g for 5 minutes.

34. BE CAREFUL!! Using suction and a 20 μL pipette tip, aspirate most of the supernatant. Since the supernatant has collected on the back of the tube, make sure you position the pipette tip so that it sucks from the front of the tube.

35. Resuspend cells in 100 μL of RLT buffer containing B-mercaptoethanol (in large cabinet under the bench). Resuspend the cells by pipetting up and down 10x and rinsing the walls carefully.

36. Vortex each eppendorf tube twice for 15 s each time to collect all the liquid at the bottom of the tube.

37. Transfer to appropriately labeled cluster tubes and freeze at -80 C.

7 Day Protocol

1. Cool centrifuge at 4°C.

2. Collect the appropriate 24 well plate from the incubator. Label 4 polysterene staining tubes corresponding to each well (A+, B+, etc).

3. Harvest cells from culture wells using a 1, 000 μL pipette by pipetting up and down to resuspend cells in the well and rinsing each well with 200 μL of staining buffer.

4. Centrifuge tubes at 300g for 5 minutes.

5. Remove cell culture supernatants. To do this, first label cluster tubes as follows: a. Specimen ID b. Date c. Supernatants- 48 hr or 7 day d. A+, B+, etc.

6. For each stimulant, using a 1000 μL pipette tip, transfer 800 μL of supernatant from the culture tube into each corresponding cluster tube. Be careful not to disturb their pellets. If you do, be sure to spin again.

7. Cap the cluster tubes and store in the -80°C freezer by the freight elevators.

8. Add 1 mL of staining buffer to each tube and vortex.

9. Wash cells at 300 g for 5 minutes at 4°C. Decant tubes.

10. Add 1 mL of staining buffer to each tube and vortex.

11. Wash cells at 300 g for 10 minutes at 4°C. Decant tubes.

12. Prepare cocktail preparation, obtaining the markers from the white box labelled "milk project" in the refrigerator in Rm. 46 near centrifuge. Be sure to store in the refrigerator (light sensitive) until ready for use.

 a.  10 μL of CD25-PCy5
 b.  5 μL CD4-PC7
 c.  5 μL of CD3-APC7
 d.  2 μL of Aqua live/dead (2 μL), which can be collected from from the refrigerator in the back of Rm 46 near the lunch room.  If you need to open a new box, be sure to dilute with 100 μL of DMSO and date and initial the tube.
 e.  1 μL of CD127-PE
 f.  Add staining buffer to reach a total volume of 50 μL per flow tube.  

13. Add 50 μL of cocktail to each tube.

14. Place in the fridge for 20-30 minutes (This could be a good time to split cells).

15. Add 3 mL of staining buffer to each tube and vortex.

16. Wash at 300 g for 10 minutes at 4°C. Decant tubes.

17. Resuspend cells in 500 μL 1x FACS lysing solution and allow to stand for 15 minutes at room temperature in the dark.

23. IN THE HOOD, prepare a solution of 20% DMSO in staining buffer. Add 2 mL of staining buffer and 500 μL of 20% STERILE DMSO to each tube. Mix gently but thoroughly.

24. Transfer the samples to appropriately-labeled cluster tubes and store samples at -80°C.


discuss this protocol


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All Medline abstracts: PubMed | HubMed