DMEM (Cellgro catalog # 10-013-CV; 4.5g/L glucose) supplemented with 10 % heat inactivated FCS. I prefer to avoid any antibiotics in my cultures, but if necessary can be supplemented to this media.

Split cells 1 à 3

Cells can be used for step-down when they are ~ 80-90% confluent. If left unattended they tend to form some clusters on top of the monolayer of cells.

Step down

Differentiation of PANC1 cells

Generally I prefer to take 80-90% confluent flasks (T-80) for step-down protocol. The passage number obtained from ATCC was passage 57 and this will be referred to herein as passage 0. The cells that were first obtained from these are passage 1 and so on. In my experience, passages 3-7 have been the best candidates in achieving decent aggregation / differentiation of these precursor PANC-1 cells. Later passages have several issues in aggregation, wherein I did see that not all the cells at later passages contributed to aggregate formation; some single cells also remained attached as typical epithelial cells even in the serum-free medium (SFM).

Media composition:

Day 0 (day of step-down):

DMEM / F12: Mixed 1:1

DMEM was obtained from Gibco (Catalog # 11885-084)

F-12 medium was from Cellgro (Catalog # 10-080-CV)

BSA (ICN Biomedicals inc. Catalog # 152401, lot 3506F): 1%

Sodium selenite (Sigma S-5261; FW 172.9): 0.0067 mg/l

Transferrin (Gibco catalog # 13008-016; 4mg/ml stock): 5.5mg/l

IGF-1 (Calbiochem catalog # 407240; 10µg/ml stock): 10ng/ml

Day 4

Day 0 media

Taurine (Sigma catalog # T-8691; FW 125.1): 0.3mM

Day 10

Day 0 media* (everything else is same except BSA increased to 1.5%)

Taurine (Sigma catalog # T-8691; FW 125.1): 3.0mM

GLP-1 (Stock 100µM): 100 nM

Nicotinamide: 1mM

NEAA: 100 µM

Procedure:

Take passage 3-7 PANC-1 cells à remove SCM and add ~ 8ml Trypsin

Dilute the trypsin digest with SFM (day 0) at the end of the digest and spin cells down

DO NOT add serum at any point to these cells once the SCM is removed and the cells are exposed to trypsin

Resuspend the pellet in day 0 SFM and pipette cells up and down at minimal pipette force for ~ 20 times.

Plate the cells on a tissue culture treated T-80 flask in 12 ml of media

Feeding cells:

I see a considerable (~10-20%) cell death after 1 day of step-down. I therefore re-feed the cells the next day with day 0 SFM and then after every 2 days starting on 4^th day with the day 4 SFM.

Re-suspend the cells ~20 times with the pipette settings set to minimal speed. This is a key step to obtaining nice tight cell aggregates.

The day 10 SFM starts on day 10 and the clusters are fed every other day. I do not spin the cells in a centrifuge at any point. I allow them to settle in a 50 ml conical tube and pull off the supernatant once the aggregates have settled down (usually in a minute or so).

ICC

Immunocytochemistry

Wash cells 3 times at room temperature (5 minutes each ) in Dulbecco’s PBS containing calcium and magnesium.

Fix cells in fresh pre-warmed (37°C) 4% paraformaldehyde for 20-30 minutes at room temperature. (Gluteraldehyde is another fixative that may be used (0.25% to 4% in 1X PBS). It cross-links proteins and generally retains their antigenicity.)

2. Wash cells once at room temperature (5 minutes each) in Dulbecco’s PBS.

3. Preps can be permeabilized at this point using chilled 50% methanol in 1X PBS. Preps should be incubated in chilled methanol for 15 minutes. Cells can also be permeabilized in 0.2% (v/v) TritonX-100 in 1X PBS. Generally this is performed for up to 5 minutes at room temperature immediately following fixation. Following permeabilization, the cells are usually washed 3X (5 min each) in 1X PBS prior to the blocking step.

4. Block for 30 minutes at room temperature in PBS blocking buffer.

5. While the cells are blocking, set up a humid chamber for performing the antibody incubations. The chamber can consist of absorbent paper, soaked in water and a plastic storage container with a lid.

6. Dilute the primary antibody (or antibodies if performing multiple labeling) in PBS blocking buffer. Generally 50-100ul of diluted antibody solution per coverslip/slide. It is a good idea to make enough primary antibody solution for one more than the number of coverslips etc. Use PAP- Pen to minimize antibody volumes.

7. Add diluted primary antibody and incubate at 4°C overnight. This generally gives lesser background. However, you may incubate the primary antibody at 37°C for 1 hour or at room temperature for 3 hours.

8. Remove primary antibody solution and wash 3X in PBS (5 minutes each wash).

9. Dilute secondary antibody in PBS blocking buffer.

10. Incubate in secondary antibody in the humid incubation chamber for 1 h at 37°C.

11. Wash coverslips / labteks as before in PBS for 3X.

Add a drop of mounting fluid; MOWIOL or Vectashield, containing Hoechst 33342. Mounting fluid should contain 10µl of (a 10mg/ml stock) Hoechst 33342 as a nuclear counterstain in 1 ml of mounting fluid. Hoechst 33342 is a nuclear stain that binds to the minor groove of DNA (absorbance max is at 340nm and emission max is at 450nm). Alternate nuclear counterstain includes DAPI (10mg/ml stock in 1X PBS), which also binds to the minor groove of DNA (359nm and emission max 461nm) and propidium Iodide (10mg/ml stock in 1X PBS) that intercalates in the DNA (absorbance max 536nm and emission max 617nm).

12. Seal with nail paint.

13. View on fluorescence microscope or place in dark at 4°C for long-term storage. Note: it is strongly recommended to obtain an image of staining ASAP; paraformaldehyde fixation is reversible with time and fluorescent signals can fade off depending on the storage and anti-fade reagents used in the mounting fluid. Therefore, one cannot be certain that the observed antigen localization is correct if it has not been viewed immediately after mounting.

To prep live mounts

Unlike fixed samples, live cells, which are adherent to coverslips, do not tolerate washes in PBS without calcium or magnesium. It is therefore suggested to supplement PBS with 0.2mM CaCl2. This is a low enough calcium concentration that will not precipitate in the presence of phosphate salts. It is obvious to state that the permeabilization step be omitted. Mounting fluid should not be added to live cell studies.

Notes:

Fixation

Paraformaldehyde should be made fresh. The use of 4% paraformaldehyde as a fixative works for most antibodies and may be used up to 10% in some cases. A non-cross-linking fixative such as cold (-20°C) methanol can be used; due to its extraction of some lipids, methanol can destroy membranous organelles. PFA fixation does not have this effect on lipids and consequently better preserves organelle structure.

Controls

Secondary Antibody Alone

In order to control for the possibility of the secondary antibody cross-reacting with cells, it is necessary to set one coverslip aside and omit primary antibody incubation step. Incubate this control coverslip in PBS Blocking Buffer only for one or more hours. This control also helps eliminate signals that are due to autofluorescence of the cells, in the case of fluorescently tagged antibodies or endogenous peroxidase and phosphatase in the case of enzyme conjugated antibodies. If the background is high, it is recommended to analyze the cells without secondary antibody as well. This will help to determine whether the background is due to the secondary antibody or the cells themselves. If the cells show a lot of autofluorescence (esp. in the green channel), then use the proper controls to take the samples over to the META for emission fingerprinting.

Preimmune/Normal IgG

In addition to omitting the primary antibody, preimmune or “normal” IgG from the animal species in which the primary antibody was raised are sometimes incubated as a control. This control rules out the potential that any signal seen with the primary antibody is actually due to nonimmune IgG cross-reacting with antigen in the target cells.

Blocking

The PBS Blocking Buffer generally should contain normal serum from the host species that is used to generate the secondary antibodies. In theory, if there are any host IgG molecules that can cross-react with the cells being studied, then the unlabeled/non conjugated IgGs present in the normal serum will cross-react during the blocking step.

For antibodies that are raised against specific peptides, it would be a good idea to block these antibodies with the specific as well as non-specific peptides so as to achieve more assuring controls. To block antibodies with peptides, use 4X by weight, the amount of peptide to antibody.

Antibody (1mg/ml): 5 ml + Peptide (1mg/ml): 20 ml + PBS 1X: 75 ml à Incubate O/N on a rotator at 4°C. Next day spin @14K for 2’. Remove the top 95 ml, further dilute to appropriate working concentration and proceed for immunohistochemistry.

Solutions

Dulbecco’s PBS

139mM NaCl

2.7mM KCl

0.75mM CaCl2

0.48mM MgCl2-6H20

8.8mM Na2HPO4-H2O

1.48mM KH2PO4

PBS (without calcium and magnesium)
4% paraformaldehyde

Dissolve paraformaldehyde in pre-warmed Dulbecco’s PBS and then add one or two drops of 10N NaOH.

Blocking Buffer

4% normal donkey serum in PBS

note: 1-4% BSA (Fraction V) may also be used.

Recipe for MOWIOL

Tris 0.2 M, pH 8.5 (MW: 121.14) prepared by adding 580 mg Tris to 24 ml water.
Add 12 g of glycerol
Add 12 ml of dH₂O
Stir on a hotplate at no more than 55°C
Add 4.8g of PVA in small batches and stir till entire contents dissolve
Then add 2.5% (w/v) of DABCO
Cool to RT
Aliquot 1 ml into eppendorf tubes.
Freeze at –20 °C and store in dark.

Hoechst (345/478) may be added prior to use at 10-20mg / ml

Transwell

Cell migration: Transwells

Place desired number of transwells in conventional 24 well plate (I generally attempt to do 15 to a maximum of 18 at a time). Trypsinize PANC-1 cells (passage 4-8) and aliquot out 30,000 cells/½ ml/well. Put cells in DMEM containing 10% FCS and place in top of transwell. In addition to the transwells, plate 30,000 cells/½ ml in 24 well dish (as a control for trypsinization). * Shake the plate sideways after putting it in the incubator so that cells are evenly distributed on the transwell membrane and do not accumulate in the center (or edges).

Let cells attach for around 2 to 3 hours at 37°C.

1) In a new 24 well plate, place 400 µl media (minimal media with 0.05% gelatin (stock 2%) with appropriate potential migration factor(s) in bottom wells (in triplicates)). (Use 1000 µl BT tip to transfer media and don’t empty the tip completely so there are no bubbles introduced in the bottom well). Use appropriate controls:

positive control and minimal media with .05% gelatin alone

2) Remove media from cells that have been growing in transwells using 10µl BT tip. Trypsinize cells with 500 µl trypsin until cells round up (use control wells to determine this). Remove trypsin with 1000 ul BT tip. Move transwells to wells with potential migration factor(s) in 24-well plate. Move all transwells. Add 100 ul minimal media and 0.05% gelatin to upper transwell using 100 ul BT tip (take care that you do not introduce bubbles). Put membrans into well on a slate to prevent bubbles.

3) Make sure there are no bubbles under membrane!!!!!!

4) Incubate for 3 ½ hours.

5) Move transwell to an empty space in the 24-well plate.

5) Remove media from top of membrane and wipe top (only) of membrane with a cotton Q-tip (use ~4 Q-tips/membrane).

6) Start fixing cells. After that count cells in the bottom of the 24 well dish (wells that the membranes sat in during the 3 ½ hour incubation.

To fix cells

1) 4% paraformaldehyde in dulbeccos PBS- pH 7.4 ( 1 g paraformaldehyde in 25 ml DPBS with 2-3 drops of 10n NaOH - heat to 60C for a while, adjust pH to 7.4, use fresh) to top and bottom of transwell for 10 minutes.

2) Count cells in bottom of transwell.

3) Suck off paraformaldehyde in membranes and on bottom of well.

4) Add syto-61 in PBS with Ca/Mg (1/1000- stock is 5 mM, working solution is 5 uM, - keep in dark)(freezer storage box # 6 or 7), approx 400 ul in bottom of transwell, 100 ul top, 37°C X 20 minutes (Syto-61 is kept in freezer, stains nucleus). (Keep this stain in the dark). Also add 20 ul stock PI (10 mg/ml in DPBS)/10 ml of PBS.

5) Suck out dye- rinse c PBS once (Fill transwells with PBS from top and let drain down).

6) Using plain glass slides. Cut membrane with razor blade and put topside down, 2 membranes/slide, then put 1 or two drops vectashield on membrane add one cover slip for 2 membranes and seal with nail polish.

7) Count whole membrane under 10X objective (approximately 15 fields), use DS red filter (Rhodamine).

Original reference is Klepes, V.E., et al, Journal of Cell Science 114, 4185-5195 (2001).

Minimal Media for PANC-1 cells

For 100 ml of media

DMEM (low glucose) + F12 (1:1)

ITS (Gibco # 51300-044) (fridge)……………1000 ul

NIC (100 mM stock, 10 mM final)…1ml, fw =122.1 (1.2g/100 ml=.1M) or add powder directly to media 0.122g/100 ml

KGF (1000X) (10 ng/ul stock)………100 ul

BSA fatty acid free(1% final)(ICN # 152401) (fridge)…………..1.0 g

islet isolation

Mouse islet isolation

Euthanize mice with compressed CO₂ followed by cervical dislocation to ensure any discomfort to the animal. Submerge in 70% EtOH to disinfect. All of the following procedure, unless otherwise specified, is performed in asceptic conditions on ice.
Open the mouse to expose the spleen.
Pull out the spleen and the majority (~70%) of the pancreas attached to the spleen. (You may also pull out the mesenteric mass of the pancreas at this time).
Remove pancreas and put them directly into 20ml DMEM, in a 50 ml conical tube. Pool the pancreas from 3-6 mice. At this time make sure that there is no fat attached to the excised tissue (fat usually floats) and wash the tissue with cold DMEM medium for a couple of times.
Remove as much media as possible and transfer pancreas to a 10cm dish.
With scissors chop tissue to pieces around 2-5 mm3.
Transfer tissue to 7ml collagenase (3mg/ml) in a 15ml tube (this is generally enough for pancreas from 4-5 mice).
Seal tube with parafilm and shake vigorously in a 37˚C water bath for about 3-5min, continuing to shake gently for another 3-5 minutes or so or until the tissue is digested (no large visible chunks).
Add 2-3 ml FCS to inactivate Collagenase.
Spin For 3 min at 150xg.
Wash 2 x in RPMI.
Plate cells in non-tissue culture treated flasks in RPMI (containing 10% FBS).
Incubate at 37˚C for 3 days in RPMI (May need to Refeed inspect for exocrine and ductal cell attachment and death).
Spin down islet at 150xg for 3min.
Resuspend in CMRL 1066 + 10%FCS + Glutamine + penn/strep.
Plate islets on tissue culture treated dishes and maintain cells in SC CMRL.

ChIP

Chromatin immunoprecipitation

(obtained from Dr. Raghu, Mirmira, UVA)

Reagents:

ChIP sonication Buffer (1% Triton X-100, 0.1% Deoxycholate, 50 mM Tris 8.1, 150 mM NaCl, 5 mM EDTA):

10 ml 10% Triton X-100
1 ml 10% Deoxycholate
5 ml 1 M Tris-Cl pH 8.1
1 ml 0.5 M EDTA
3 ml 5 M NaCl
80 ml Water
Just before use, add 10 ul Aprotinin, 10 ul Leupeptin, and 5 ul PMSF to each 10 ml.

High Salt Wash Buffer (1% Triton X-100, 0.1% Deoxycholate, 50 mM Tris-8.1, 500 mM NaCl, 5 mM EDTA)
10 ml 10% Triton X-100
1 ml 10% Deoxycholate
5 ml 1M Tris-8.1
1 ml 0.5M EDTA
10 ml 5M NaCl
73 ml Water

LiCl Immune Complex Wash Buffer
25 ml 1M LiCl
5 ml 10% IGEPAL
5 ml 10% Deoxycholate
1 ml 1M Tris-8.1
200 ul 0.5M EDTA
64 ml Water

Protease inhibitors (add 10 ul of each to 10 ml of PBS or sonication buffer)
Leupeptin 2 mg/ml in water
Aprotinin 2 mg/ml in water
PMSF 0.2 M

5 M NaCl

1X TE Buffer (10mM Tris, 8.1, 1 mM EDTA)

0.5 M EDTA

1 M Tris-Cl, pH 6.8

Protein A/G Agarose (Santa Cruz)

Proteinase K (19 mg/ml, Boehringer Mannheim # 1964372)

10X proteinase K buffer

Elution Buffer (1% SDS, 0.1 M NaHCO3, 0.01 mg/ml Herring sperm DNA, 2ng/ml CMV beta Gal plasmid)

10 mg/ml Herring Sperm DNA

37% Formaldehyde (ACS reagent grade)

1.25 M glycine

Protocol: Generalized for many cell types, but may require optimization for specific cell types (tested with mPAC, bTC3, aTC, HEK, HeLa, NIH3T3).

For all the following steps, use the pipets that are specifically designated for ChIP use only.

Day 1:

1. To each 10 cm dish of cells, wash plate once with 10ml of PBS, then add 10 ml of Fresh PBS and add 270 ul of 37% formaldehyde, swirl gently to mix, and place at room temp 10 min.

2. At the end of the incubation, add 1 ml of 1.25 M glycine, swirl to mix.

3. Aspirate medium

4. wash plate with 10 ml cold PBS x 2. Aspirate PBS completely after the second wash
.

5. add 500 ul of cold PBS + protease inhibitors and scrape cells, collect in a 1.5 ml centrifuge tube. At this point you should pool three plates worth of cells together in the same tube (I suggest using a 2 ml siliconized eppendorf tubes for this purpose).

6. centrifuge at 2000 rpm for 2 min at 4 º C.

7. remove and discard PBS

8. add 600 ul of ChIP sonication buffer + protease inhibitors, and resuspend pellet (you can vortex vigorously at this point).

9. Place on ice for 10 min
.

10. Sonicate: We use a Misonix Sonicator (model S-300 sonicator with 2.5 in diameter cup horn and 8-place sample holder for sonicating multiple samples). The cup horn should be filled with an ice-water mixture. We recommend the following sonication settings:

Amplitude setting: 4

15 five second pulses with 15 second cool-down intervals between pulses. The shearing may be more efficient if the tubes can be placed at a 30-45 degree angle.

Alternatively, you can use a setting of 4 and deliver two 90 sec pulses with 1 min 45 s cool-down interval between pulses.

We found that the two protocols are very similar in shearing DNA to the 500-2000 bp range, however, the first protocol minimizes heat build-up.

11. Centrifuge at maximal setting at 4 C for 10-15 min.

12. Remove the supernatant into a fresh siliconized 1.5 ml eppendorf tube. This is the Whole Cell Extract (WCE), and can be stored at –80 C at this point, if desired.

13. Check protein levels by making a 1:10 dilution of a sample of the extract in water and doing a Bradford assay. Use 100 ug of WCE per antibody immunoprecipitation. If not enough extract is available, bring the solution up to 100 ug protein by adding purified/acetylated BSA.

14. Pipet 100 ug WCE into a fresh siliconized tube containing ChIP sonication buffer with protease inhibitors to a final volume of 1 ml.

15. Pipet 10% input into a 1.5 ml eppendorf tube (not siliconized, because it will dry faster after 70% EtOH wash—see later), and store at -20 C—you will need it later.

16. Add appropriate antibody volumes to each sample:

5 ul for histone antibodies from Upstate (acH3, acH4, 2meK4, 3meK4)

10 ul for histone antibody 1meK4

5 ul for Pdx-1 rabbit polyclonal antiserum, RNA polymerase antibody from Santa Cruz

2 ul for CTD RNA polymerase Ab

10 ul for RNA polymerase Ser2 and Ser5 Ab

17. Place the samples on a nutator in the cold room, and incubate overnight.

Day 2:

18. Resuspend Protein A agarose so that it forms a uniform suspension. Using a pipet tip with the end clipped off, add 45 ul of this suspension to each immunoprecipitation. Resuspend the protein A agarose each time before adding to the next sample, as it settles quickly.

19. Add 2 ul of a 10 mg/ml solution of herring sperm DNA

20. Place back on the nutator at 4 C for 1-2 h.

21. Centrifuge the samples at 4 C for 2 min at 2600 rpm.

22. Carefully remove the supernatant using a P-1000 and place it in a tube and label it “sample X—sup.” Place this at –20 C in case you need it later.

23. Add 1 ml of COLD ChIP buffer (no protease inhibitors), invert the sample to resuspend the resin, and centrifuge for 2 min. at 2600 rpm.

24.remove and discard the supernatant.

25. Wash X1 with High Salt buffer, X1 with LiCl buffer, and X1 with TE

26. Add 250 ul of Elution buffer to the resin, and place on a nutator at room temp. for 15-20 min.

27. Centrifuge at top speed to pellet the resin, remove the supernatant to a fresh tube.

28.Repeat the elution step (step 26), and combine the supernatants. (you can now discard the tube containing the resin pellet)

29. At this time, add 500 ul of elution buffer to the “10% input samples” from step 15.

30. Add 20 ul of 5 M NaCl to each sample, vortex to mix, and place in a 65 C bath for 3-4 h.

31. Add 1 ml of ROOM TEMP ethanol to each sample place at –20 C overnight.

Day 3:

32. Next day, spin the samples at top speed at 4 C for 15-20 min. to pellet the precipitated protein/DNA.

33. Aspirate off the supernatant, add 1 ml of ice cold 70% ethanol, spin again at 4 C for 5 min.

34. Aspirate off the sup, allow to air dry for 5-10 min.

35. Dissolve the pellet in 100 ul of TE.

36. Add 11 ul of 10X Proteinase K buffer, and 1 ul of a 19 mg/ml proteinase K solution.

37. Add 390 ul TE

38. Add 500 ul of PCIAA (phenol/chloroform/isoamyl alcohol)

39. Vortex for 30-60 sec and spin at max speed for 1 min.

40. Carefully remove the aqueous phase and transfer to a new tube.

41. Add 20 ul of 5 M NaCl, vortex to mix

42. Add 1 ml of room temp 100% EtOH

43. Incubate at -20 C overnight or -80 C for 1 h

44. Centrifuge at max speed for 20 min. at 4 C

45. Remove supernatant and add 1 ml of cold 70% EtOH

46. Centrifuge at max speed for 5 min at 4 C

47. Air dry tube

48.Resuspend in 100 ul of TE (Note: you may NOT “see” a pellet, but that’s OK!)

49. You’re ready for PCR.

Cell counting and basics