methods

METHODS FOR CELL CULTURE

LAB PROTOCOLS AND STANDARD OPERATING PROCEDURES FOR INNOVABONE PROJECT

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Table of Contents

1.0 Osteoblast Isolation from Mouse (English)2.0 Osteoclast isolation from Mouse (German)3.0 Osteoclast isolation from Rabbit (English)4.0 Procedure for generating mouse OB/OC co-culture (English)5.0 Standard operating procedure for splitting adherant cells (Eng)

5.1. Table: Medium formulations by cell and assay type (Eng)6.0 Standard Operating Procedure for freezing cells (English)7.0 Standard operating procedure for thawing cells (Eng)8.0 Standard operating procedures for cell counting

8.1. Using the heamocytometre (Eng)8.2. Using Casey (German)

9.0 Preparation of disks and coverglasses for cell culture10.0 Seeding cells for co-culture 11.0 Osteoblast Assays

11.1. Alkaline Phosphotase Assay (English)11.2. Mineralisation Assay (English)11.3. Coculture fixation procedure (English) 11.4. Actinium/DAPI stainging (English)

12.0 Osteoclast Assays12.1. Pit Assay (German + English)12.2. Immunohistochemistry osteoclast protocol (German +

English)12.3. Actin Ring 12.4. Apoptosis12.5.

13.0 Preparation of standard solutions 13.1. Trypsin

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13.2. PBS14.0 Other something15.0 Other something

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1.0 Osteoblast Isolation from Mouse Calvaria

Preparation of the enzyme solution(-20C) Collagenase 0.1% 0.10g 0.05g(+4C) Dispase 0.2% 0.20g 0.10g

-mem 0% 100ml 50ml

wearing a face mask, weigh, mix and filter sterilize

Preparation of the 10x Tripsin EDTA solutionEDTA 0.155g Trypsin 0.50gNaCl 0.90g

dH2O up to 100ml

First add the Trypsin to the most of the water, and adjust the pH with NaOH to 7.7Then add the Trypsin ad the NaCl, then adjust the pH to between 7.4-7.6Wearing a face mask, weigh, mix, filter sterilize and aliquot. Store at -20C

Make 1x PBS-TC,(without Ca/Mg) from the 10x PBS by diluting (adjust the pH =7.2)

1. Cut the head off 1 day old mice and place in a petri dish with cold PBS2. Isolate the calvaria from the head. (50-60 calvaria is good for seeding in 6

petri dishes)3. Place the calvaria into a centrifuge tube and add 8ml of the enzyme solution4. Place at 37C for 10 mins, in the shaking incubator at 200 cycles/min5. Transfer the solution with a transfer pipette into a new tube, and place the

tube on ice. This is fraction 1 which is not needed but saved till the end of the isolation.

6. Add a further 8-10 of the enzyme solution to the calvaria and place in the shaking incubator for another 10 minutes

7. Remove the enzyme solution into a new tube and keep on ice (this is fraction 2)

8. repeat steps 6 and 7 twice more, each time adding the fractions 3 and 4 to fraction 2.

9. Reapeat a final time using 9ml of the enzyme solution, and placing for 20 minutes in the shaking incubator.

10. This fraction 5 is also added to the previous fractions (fractions 2,3,&4).11. Centrifuge at 4C 1500 rpm, 5 minutes12. Remove the supernatant and resuspend the pellet.13. Add 5 ml of the media (-mem with 10% FCS &1% P/S) and disperse, add

5ml further and again disperse14. With the same media, fill to 48ml

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15. Using 6 petri dishes, add 8ml of the cells to each petri dish and add a further 2ml media. These volumes can be altered to ensure that there is enough for all the petri dishes that you have.

16. Incubate for 24hours17. Change the medium (this will remove most other cell types that will be

present ‘contaminating’ the OB culture)18. Incubate for 24 hours19. Dilute the 10x trypsin with PBS to get a 1x trypsin solution ( you need about

2ml per petri dish) – when splitting plan on making 4-5 new petri dishes for every existing petri dish. Ie split 1:5 if you are leaving over the weekend, or 1:4 if it is during the week.

20. Remove the media, add 5ml of warm PBS (37C), and wash gently and remove

21. Add 2ml of the 1x trypsin (37C) and place in the incubator till the cells begin to lift. Use a wide mouth pipette to collect the cells an place in the centrifuge tube

22. Add media to the plate using a plastic (wider mouth) pipette, wash the plate using a small volume each time, adding the solution to the centrifuge tube

23. The total volume of media after washing should be 10ml in the centrifuge tube

24. Add 5ml of cold media to the petri dish, up and down three times to wash, transfer this to the next petri dish and again up and down. Use this same solution to wash all petri dishes and then transfer to the centrifuge tube. (this cold media removes lifts any remaining cells)

25. Repeat point 24 again26. Centrifuge at 4C 1500rpm for 5 mins27. Repeat points 12 to 1428. This time put 2ml of the cells into each petri dish, with 8mls of media. (this

volume can be adjusted depending on the splitting ratio desired)29. Incubate for 48 hours30. The freezing media is made as 20% DMSO in -mem (with 0%) – filter

sterilize (make at 4C don’t warm)31. Repeat steps 20 to 26. Use 1 centrifuge tubes for ever 6 Petri dishes32. Remove media, resuspend the dry pellet and disperse in 5ml of the 10%

media. Combine all the tubes. 33. Count the cells: take approximately 50

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2.0 Osteoclast isolation from Mouse (Bone marrow cells)

Euthanise mice and isolate the 2 posterior feet, and add to sterile PBS or a-MEM using aseptic technique, Isolate femurs in a-MEM and flush out bone marrow using G23 hypodermic needle with 1 ml syringe in Petri dishes.Homogenize bone marrow and Collect into 50 ml falconCentrifuge at 1500rpm, 4°, 5 min.Discard supernatant and disperse pellet in 10% a-MEM (non heat inactivated).Remove incubated medium from OB and seed bone marrow cells on OB + Vit D and PGE2Incubate at 90% RH, 5% CO2, 37°cchange medium after 2 days.Time point fixation: day 4-5.

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3.0 Osteoclast isolation from Rabbit

Medium: Alpha MEM = 218 270 µl, FCS hi 5% = 11500 µl, P/S 1% = 230 µl

Protocol for 1 rabbit:

Prepare and set up under the laminar airflow chamber: 3x 50 ml-centrifuge tubes, one of those cut at 35 ml Box with ice 2x big Petri dishes: 1x with sterile PBS and 1x with Medium Plastic pipette Scissors, Scalpel and pinzette in alcohol 10 ml plastic pipette with a big hole (in order not to damage cells) Start the centrifuge at 600 rpm, 5 min, 4 °C

Put the legs of the rabbit in the Petri-dish, remove all muscles with the scalpel and scissors.Put in another Petri dish with sterile PBS. Under the laminar cut the clean bones into pieces

1. Take a 50ml tube already cut to 35 ml, put 6ml medium inside and add bones pieces.2. Take the scissors out from alcohol and cut for 2 minutes the bone.3. Add 7 ml 5% medium so that the cut legs are in 13 ml medium4. Take supernatant &transfer with plastic transfer pipette in another 50ml tube on ice

Point 2-4 you repeat 2 times. (total we cut 3 times)

Add 13 ml of 5% medium, in 3 steps (5ml, 2x 4ml) to the bone, and transfer the medium with bones in a 50 ml centrifuge tube - Vortex 30 seconds

Wait 2 minutes till bones settle, take the supernatant and add to the other supernatant on ice (now we had put 4 times the supernatant in the tube)

Put the tube in the centrifuge: 5 minutes centrifuge 600 rpm, 4°C Remove the supernatant and take the pellet

Important: dont use glass pipette for OC, but plastic pipettes with big opening in the tip.

Shake the centrifuge tube with the pellet. Add 5 ml medium and MIX very well Add 12 ml to have a final volume of 17 ml. (Add like this: 5 ml + 5 ml + 7 ml,

and mix every time) Seed 300 µl of cells in each well (48 well plate), or put 100 l Cell suspension

on one glass, which lies on sterile Para-film in a Petri-dish 3 hours of incubation Remove the medium with the pump, carefully, put 500 µl 5% medium back in

the wells, remove the medium again Add 500 µl 5% medium (with treatments when there is the need), or put the

glass into the well-plate and add the medium. 48 hours incubation

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4.0 Procedure for generating co-cultureSeed at OB in a-MEM (heat inactivated) at least 6 hours before addition of OC.Remove incubated medium from OB and seed bone marrow cells freshly obtained as per OC isolation protocol, on OB + Vit D and PGE2Incubate at 90% RH, 5% CO2, 37°cMedium change after 2 days.Time point fixation: day 4-5

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5.0 Standard operating procedure for splitting adherant cells

5.1. cell lines in flasks1. All procedures should be conducted in a sterile tissue culture

hood.

2. Prepare appropriate media with 10% FBS and appropriate supplements for the cell line.

3. Prepare a new flask with complete media, ensure media covers surface of the flask (approximately 10ml for a medium size flask). Place in the 37 °C incubator to equilibrate, until flask is required.

4. Remove old media from cells in tissue culture flask.

5. Add trypsin/EDTA to the cells after you have removed the old growth media. I use about 2ml of the trypsin/EDTA solution on a medium size tissue culture flask. Swill the solution around for 3-4 seconds very gently and siphon away the trypsin/EDTA quickly.

6. Add 2-3ml trypsin/EDTA for 3-10mins, until the cells come away from the flask, store in the 37°C incubator during this time

7. Add 5-8 ml of the media to the cells from the first tissue culture flask and removed the cells from the flask using a pipette, and place in a 15ml eppendorf tube.

8. Spin in a centrifuge at 1.2rpm for 3minutes. A cell pellet should then be visible at the bottom of the tube. Tip off excess solution, and re-suspend the cell pellet in about 10ml of media.

9. If necessary count the cells using a hemacytometer using SOP,

10. Dilute cells to the appropriate number for experiment, or for the number required for seeding into a new flask. Usually for propagation a few drops to 1ml is appropriate depending on the cell pellet size

11. Add a few drops -1ml of the cells to the pre-prepared flask and store in the incubator.

12. This should be repeated before cells reach confluence (at a maximum of about 70% confluence)

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5.2. Splitting of Primary OB

1. Prepare warm sterile PB S w/o Ca and Mg.2. Prepare Trypsin 1x (final concentration 0.2 5% from Boehringer Ingelheim

and 0.05 % from our lab.) with warm sterile PB S w/o Ca and Mg.3. Remove incubated medium from petri dishes.4. Wash twice with warmed sterile PB S w/o Ca and Mg.5. Add Trypsin solution to cells (4 ml/10 cm Petri dishes, 200 µl/ 6

wells/plate, and 100 µl/12 wells/plate).6. View the detachment of cells under the microscope, to ensure that cells

have lifted.7. Stop Trypsin activity by adding an amount and 10% a-MEM (10 ml/10 cm

Petri dishes, 2ml/ 6 wells/plate, and 1ml/12 wells/plate).

5.3. Table: Medium formulations by cell and assay type Cell Assay Medium FCS P/S SupplementsPrimary OC mouse

Isolation, seeding and splitting

-MEM 10% 1%

Primary OC rabbit

Isolation, seeding and splitting

-MEM 1%

Primary OB mouse

Isolation, seeding and splitting

-MEM 10% 1%

Primary OB mouse

ALP/ Mineralisation

-MEM 10% 1%

OC/OC Coculture -MEM 1% + Vit D and PGE2RAW 264.7 Seeding and

splitting1%

Mg63 Seeding and splitting

1%

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6.0 Standard operating procedure for freezing cells

13. All procedures should be conducted in a sterile tissue culture hood.

14. Prepare appropriate media with 10% FBS and appropriate supplements for the cell line.

15. Remove old media from cells in tissue culture flask.

16. Add trypsin/EDTA to the cells after you have removed the old growth media. I use about 2ml of the trypsin/EDTA solution on a medium size tissue culture flask. Swill the solution around for 3-4 seconds very gently and siphon away the trypsin/EDTA quickly.

17. Add 2-3ml trypsin/EDTA for 3-10mins, until the cells come away from the flask, store in the 37°C incubator during this time

18. Make a 50:50 mixture of the FBS:DMSO, Aliquot 100 µl of solution into cryo-vials

19. Add 8 ml of the media to the cells from the first tissue culture flask and removed the cells from the flask using a pipette, and place in a 15ml eppendorf tube.

20. Spin in a centrifuge at 1.2rpm for 3minutes. A cell pellet should then be visible at the bottom of the tube. Tip off excess solution, and re-suspend the cell pellet in a minimum of media (cryoflask number x 1ml).

21. If necessary count the cells using a hemacytometer, so that a minimum of 500,000 cells is in each vial

22. Add 900ul of the cell/media mix to the DMSO in the cryovials.

23. Add the vials to the “Mr. Frosty” container as per the SOP

24. Store the vials in the “Mr. Frosty” at –80°C for a minimum 8 hours.

25. Transfer vials to the liquid nitrogen

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26. Following one week and thaw one of the vials you deposited in the liquid nitrogen. This allows one to verify the viability of the cells and to eliminate the possibility of contamination.

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7.0 Standard operating procedure for thawing cells8.0 Standard operating procedures for cell counting

8.1. Using the heamocytometre8.2. Using Casey

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9.0 Preparation of disks and coverglasses for cell culture9.1. Disks

1. Must be done at least 48 hours prior to the addition of cells!2. Place each sterile disk in a well of a 24 well plate, under the laminar

hood. 3. Place 500µL of the same media (with FCS and %PS) that will be used

for cell culture into each well. 4. Incubate for a minimum of 48 hours at 37°C

5. Disks and porous materials absorb the media, and thus if this is done for less than 48 hours then the media with cells will all be absorbed and the experiment will not be possible.

9.2. Cover glasses (coverslips)1. Presterilise the coverslips by leaving in alcohol overnight. 2. The coverslips are then dried by tapping on paper, and placed in a

metal box lined with a piece of paper towel. Do not stack lay out flat in the box. A second paper towel is placed over the top.

3. Place sterilation tape on the box, and heat sterilise for 3 hours. 4. These can be stored untill use taking care not to tip the box, so that

the coverslips stay layed out nicely. 5. Open the sterile box under the laminar hood. Place up to 8 coverslips

into one petri dish for preincubation. Lay out flat. Add 10 ml of the appropriate media to each of the petri dishes.

6. Leave in the incuabtor at 37°C for 24 hours. 7. Then transfered to 6 well plates for the experiment.8. To transfer, first drip dry the coverslips for a few seconds, tap on a

paper towel gently and place in the 6 well plate, for the addition of the cells

9. The preincubation must not be done in the 6 wel plates directly. If it is done directly in the 6 well plate then the cells will not only stay on the coverslip, but also run off into the plate because it has been wet.

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10.0 Seeding cells for co-culture11.0 Osteoblast Assays

11.1. Alkaline Phosphotase Assay

• Preparation of the culture medium:α- MEM, 10 % FCS h.i, 1 % P/S Mix well,

Preparation of the mineralization medium:

1 % VIT C 100x,1 % βGP 100x, 10 % FCS h.i, 1 % P/S in α-MEM (subtract all volumes, and make up to 100% with a-MEM)

The vitamin C is unstable and therefore the medium has to be prepared justbefore use. The mineralization medium has to be approximately 37°C before it is added to the cells. Warm up the mineralization medium in a water bath beforeuse.

Preparation of the βGP-solution: 10 mM in PBS.MW βGP = 216 g/molStock solution: 100x.2,16 g in 10 ml PBS10,8 g in 50 ml PBS

Preparation of the lysisbufferDissolve 0,05 g Triton in 98 ml PBS without Mg2+/Ca2+Add 2 ml 1M HEPESThe total amount volume is 100 ml and is it stored in room temperature

Preparation of the standard solution for ALP assay:The standard solution contains 4-nitrophenol and it is stored in the refrigeratorCalculate how much standard solution is needed:Need 1,26 ml for one row with standards – always make duplicates

Examples:Dilute 25 µl standard solution in 5 ml 0,02 N NaOHDilute 20 µl standard solution in 4 ml 0,02 N NaOHDilute 15 µl standard solution in 3 ml 0,02 N NaOH

Preparation of ALP substrate:The substrate is 40 mg p-nitrophenylphosphatedisodium powder, which issolved in 10 ml purified water. The powder is stored at -20 °C. Transfer 1 ml into10 eppendorfs and store at -20 °C.

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Preparation of A´:Ratio: 20 µl S + 1000 µl ACalculate how much needed, need 25 µl for each well.Example: 1700 µl A = 1700 µl/1000 µl * 20 = 34 µl SReagents is in the protein assay kit

Preparation of the Alizarin Red-S 40 mM:Prepare 50 mlMw = 342,3 g/moln(ARS) = C * V = 40 mM * 0,05 L = 2 mmolm(ARS) = n * Mw = 0,002 mol * 342,3 g/mol = 0,6846 g

0.002 molWeigh in 0,6846 g Alizarin Red-S and dissolve in 50 ml waterAdjust the pH to 4,2 with a pH-meter

•

Preparation of the Alizarin Red-S 4 mM:Prepare from ARS 40 mMNeed 350 µl ARS 4 mM, prepare 500 µl for safeC1V1=C2V2

40 mM * V1 = 4 mM * 500 µlV1 = 50 µlWater needed: 500 µ - 50 µl = 450 µlDilute the 50 µl of ARS 40 mM in 450 µl water

Preparation of the standard solution for making the standards in themineralization assay:Need 3052,5 µl, prepare 3500 for safe350 µl ARS 4 mM + 3150 µl 10 % CPC = 3500 µl standard solution

Preparation of the 10 % CPC solution:10 g cetylpyridiniumchloride in measuring flask with 10 mMsodiumphosphatebuffer, pH 7 filled up to 100 ml.Place the solution in water bath

Preparation of the phosphatebufferSolution A 0,2 M: Dissolve 6,81 g KH2PO4 in 250 ml waterSolution B 0,2 M: Dissolve 7,05 Na2PO4 in 250 ml waterPhosphatebuffer pH 7:

19,5 ml solution A30,5 ml solution BmilliQ-water to 100 mlPhosphatebuffer pH 7, 10 mMDilute the phosphatebuffer, pH 7 in water 1:10 and set the pH to the exact valueof 7,0

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Seeding of osteoblasts4 plates -12 wells, 1 ml/wellFor each plate we need 2 million osteoblasts (OB). Calculate how much OBneeded and see, which vials that are available. For 4 plates, one needs 8 millionof OBs.The vials are frozen on liquid nitrogen.Prepare LAF-bench, switch on water bathIf there is a substance needed that is stored in the freezer, get it now for thawingNeed 48 ml culture medium. Prepare more for medium changes later, calculatehow much needed.

Mark plates with name, date and cell typeAdd 0,5 ml of the culture medium in each well – do not need warm mediumTransfer 5 ml of the culture medium to a tubeGet the OB from the liquid nitrogen; remember polystyrene to carry it inThaw the vials of osteoblasts in a water bath (37 °C)Swab the vials with 70 % alcohol and transfer the contents of the vials to thetube with a plastic pipetteWash the vials once and mix carefully with the culture mediumFill the tube with 5-10 ml at a time, up to 25 ml and mix carefully between, (forsafe, one extra ml) and disperse and dilute the osteoblasts so that the number ofOB is 2 millions for 12 wells.Add 0,5 ml of the mix with osteoblasts in each well.Check the cells in the plates in the microscope before incubation at 37 °C.Day 1-5Control the confluence of the osteoblasts under a microscope.Change medium to the mineralization medium if the cells are confluent

If treatment is going to be added, it is together with the mineralization medium,

when the osteoblasts are confluent

Medium change:Calculate how much medium needed, 1 ml/well. Prepare 1 well extra for safe.Put the medium on water bath before adding it to the cells ( this is just up to p6 of the doc keep copying)

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11.2. Mineralisation Assay

Preparation of the mineralization medium:

Calculate how much needed, example 50 ml:

1 % VIT C 100x 0,5 ml (500 l)μ1 % GP 100x β 0,5 ml (500 l)μ-MEM, 10 % FCS h.i, 1 % P/S 50 ml – 0,5 ml – 0,5 ml = 49 ml

The vitamin C is unstable and therefore the medium has to be prepared just before

use. The mineralization medium has to be approximately 37°C before it is added to

the cells. Warm up the mineralization medium in a water bath before use.

Preparation of the Alizarin Red-S 4 mM:

Prepare from ARS 40 mM stock solution by 1:10 dilution

For a 12 well plate prepare 500µL of ARS 4 mM

So dilute the 50 l of ARS 40 mM in 450 l water

Preparation of the standard solution for making the standards in the

mineralization assay:

For a 12 well plate prepare 3500µL (need 3200)

350 l ARS 4 mM + 3150 l 10 % CPC = 3500 l standard solution

Preparation of the 10 % CPC solution:

10 g cetylpyridiniumchloride in measuring flask with 10 mM

sodiumphosphatebuffer, pH 7 filled up to 100 ml.

Place the solution in water bath

Preparation of the phosphatebuffer

Solution A 0,2 M: Dissolve 6,81 g KH2PO4 in 250 ml water

Solution B 0,2 M: Dissolve 7,05 Na2PO4 in 250 ml water

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Phosphatebuffer pH 7:

19,5 ml solution A

30,5 ml solution B

milliQ-water to 100 ml

Phosphatebuffer pH 7, 10 mM

Dilute the phosphatebuffer, pH 7 in water 1:10 and set the pH to the exact value of

7,0

Mineralization assay – day 15

To assess the amount of Ca2+ in the mineralization, we use Alizarin Red-S (ARS),

which is a coloring agent that binds calcium salts selectively.

1 mol ARS bind 2 mol calcium

Take the plates out from the incubator and remove the medium

Wash three times with 1 ml cold PBS without Mg2+/Ca2+ from the refrigerator

Fix cells with 1 ml cold ethanol 70 % for one hour in the fridge sitting on ice.

Mark eppendorf with A1, A2 and so on and while waiting: prepare the ARS 4 mM,

standard solution and standards of all concentrations

Standard solution (l) 10 % CPC (l) Concentration (µM)

15 1485 4

37,5 1462,5 10

150 1350 40

225 1275 60

375 1125 100

750 750 200

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1500 400

After 1 hour, take the cells out of the freezer and wash twice with MilliQ-water

Stain wells with

Alizarin Red-S, 40 mM, pH 4,2 and put in on a shaker for 10 minutes (RT)

6 – wells: 1 ml

12- wells: 500 l

Wash five times with 1 ml milliQ-water

Add 1 ml PBS without Mg2+/Ca2+ and put the cells on a shaker for 15 minutes

Remove the PBS, scan the plates and photograph

Settings for the scanner:

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Example of a scanned plate:

Decolorize with 500 l 10 % CPC solution in 10 mM phosphatebuffer for 15 minutes

on a shaker in room temperature

Transfer stains with a micropipette to the marked eppendorf and centrifuge for 5

minutes, 15000 RPM at room temperature

Add the standards (300 l/well), an empty value (300 l 10 % CPC/well) and the

stains (samples) to an ELISA- plate. Dilute stains 1:9 in the microplate (Add 30 l

sample + 270 l 10 % CPC)

Measure absorbance at 520 nm

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11.3. osteoclast fixation (inc co culture)Wash with 1 ml 37°C or room temperature PBS+Ca Mg to detach OB layer in CC or floating cells in OC culture. (500µl for bone slices and 48 well plate)Fix once for 3 min with 1 ml (500µl for bone slices and 48 well plate) of 3.7% Formaldyde at room temperature, the remove solution (this solution is prepared by diluting 37% Formaldehyde, 1:10 in PBS)Fix once for 10 min with 1 ml (500µl for bone slices and 48 well plate) 3.7% Formaldyde at room temperature.Wash twice with 1 ml ((500µl for bone slices and 48 well plate) PBS+Ca Mg at room temperature.Add 1 ml (500µl for bone slices and 48 well plate) RT PBS+Ca for storage and add sodium azide for long conservation.Store at 4 deg (cold room).

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11.4. Actinium/DAPI staining of fixed cells (English)Preparation of Reagents:

5 µl / cover slip (or per 2 disks) of „Alexa Fluor 546 phalloidin (Nr.144)“ – put in one Eppendorf- light protection!! (Put in a cupboard) – and let the solvent evaporate under N2 gas) – do up to 10 coverslips per time

wrap the Eppendorf with aluminium foil –When it has dried out-add 200µl PBS+Ca+Mg/ into eppendorf and mix.

DAPI-dilution: 200 µl stock-solution with 10 ml PBS+Ca+Mg – light protection (2 ml is used per coverslip or per 2-3 disks)

Protocol: Prepare a metal box with parafilm attached to the bottom with

sticky tape. Put the cover slips/disks into hutches -covered with 3ml ice-cold

acetone (propanone) – 3 min (use Acetone only under the hood!!) Take the cover slips/disks out, drip dry and deposit them on

parafilm Put 200 µl of the dilution on each cover slip, 80ul on disks, add

drop by drop to different areas of the coverslip, so that the whole coverslip is covered and none runs off. – incubation for 30 min with light protection at room temperature

2x washing in PBS+Ca+Mg (fill the 6 well plate and tip upside down in the sink for coverslips- add and remove for disks)

Put the cover slips into a 6 well-plate, or 2-3 disks per well – put 2 ml DAPI-diluted solution on each well – incubate 10 min in the dark at 37°C

2-3x washing in PBS+Ca+Mg 2-3x washing in distilled water Label glass slides Put flurosave („tissue-teck“) on the glass slide (for disks use the

one in the small white bottle in the fridge) Tap the coverslip/disk edge gently on a paper towel and add the

cover slip/disk on the glass slide, with cell side on the tissue tek.- this means cell side down for coverglasses and cell side up for disks, and fix the borders with glue – drying in the dark

Storage at 4°C in a folder wrapped with aluminium foil

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11.5. something11.6. something11.7. Something

12.0 Osteoclast Assays12.1. Pit Assay

Pit Assay German and English

Isopropanol 70 % für Ultraschall, wiederverwendbar(ca 1 ml in 4ml Polypropylenröhrchen)

Pinzette in Alkohol Knochenplättchen aus Well in 70 % Isopropanol transferieren

15 min UltraschallbadZellen werden dabei entferntMit Zahnbürste restl. Zellen entfernen (betrifft Co-Kultur)

FärbungFärbungsreagenz (wiederverwendbar) 1% Toluidinblau 1% Na-Borat dest. H2O

4 min schwenken

Entfärben

Mit dest. H2O

Färbung von Humanknochen Färbung mit Hämatoxylin Mayer 1 min

(in Flaschen Sigma) können auch mit Toliudinblau gefärbt werden

(eher niedriger als 1%-ig)

English• 70% isopropanol for ultrasonication, Reusable(about 1 ml in 4 ml polypropylene tubes)• Tweezers in alcohol

1. Bone pieces are taken out of the wells and put into into 70% isopropanol using one bottle for each group2. Ultrasonicate for 15 min. Cells are removed at this time. 3. Dip each bone piece in water4. Remove remaining cells with a toothbrush (affects co-culture)

colorationColouring agent (reusable)

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• 1% toluidine blue• 1% sodium borate• made up in distilled a´waterplace in the shaking incubator for 4 mins.

decolorize by washing with dist. H2O

Staining of human bone• staining with hematoxylin Mayer 1 min(bottled Sigma)• can also be dyed with Toliudine blue(rather less than 1%)

12.2. Immunohistochistry osteoclast protocol

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Immunofluorescence for Osteoclasts

Take 2 cover slips per treatment and for controls in a 6 well plate, fixed with 3.7% formaldehyde in PBS. Control groups are always necessary, since osteoclasts normally have fluorescence.

1. Alcohol series; 40%, 70%, 80%, 90%, 100%, 90%, 80%, 70%, 40%.a. Put 2ml in each well of the 6 well plate. Swirl the solution briefly and

then remove.b. This dehydration is necessary to make the cell wall permeable to the

antibody.

2. Solution 1. 1ml per wella. Prewashing will improve the surface wettability.b. For approximately 24 coverslips you need 100ml PBS with

Ca2+/Mg2+c. Add 100mg 0.1% BSA – kept in the fridged. Add 50mg 0.05% Saponin (from sigma) – kept in the chemical

cupboarde. Dissolve for a long time on a magnetic stirrer.

3. Add 2ml of PBS with Ca2+/Mg2+ to each well, and wash 2-3 times4. Solution 1 add 1ml to each well. Taking a break at this step is possible since

the cells are fixed.

5. Put Coverslips on parafilm a. Add 100µL of Solution 2 b. To prepare enough solution 2 for 12 coverslips, add 65µL of NGS (5%)

to 1235µ@ of solution 1. This makes a total of 1300µL (ratio is 1:20)c. Nb. NGS is natural goat serum (Ziegenserum) – kept in shelf 2 of the

freezer at -20°C.d. Stand for 30 minutes covered

6. Dry with kitchenpaper towel and put coverslips on new parafilma. Add 100µL of solution 3 (primary antibody)b. Stand for 2 hours covered.

Instructions to prepare solution 3- primary antibody. Primary polyclonal antibody directed against phosphorylated tyrosine residues. Antiphosphorylated rabbit in 50% glycerin solution, since this has the glycerin then it will not be solidified even though it is in the freezer. It is kept in the electrophoresis room at -20 in shelf 1.

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For 12 coverslips mix in an eppendorf; 1,290µL of solution 1 +65µL of NGS (5%) + 26µL of primary Antibody (3%) (the ratio is 1:50) total volume is 1300µL, collect the contents of the eppendorf by a brief centrifugation.

7. Dry, and wash with solution 1, three times in a 6 well plate.8. Put the coverslips on new parafilm

a. add 100µL of solution 4b. wrap plate in Aluminium foil and stand for 1hr

To make solution 4 the fluorescent antibody- rabbit (secondary antibody) – the antibody is kept in the fridge. It is light sensitive and so the eppendorf should always be wrapped in Aluminium foil and stored in the dark.

For 12 coverslips mix in an eppendorf; 1,290µL of solution 1 +65µL of NGS (5%) + 26µL of secondary Antibody (3%) (the ratio is 1:50) total volume is 1300µL, collect the contents of the eppendorf by a brief centrifugation.

9. Wash in the 6 well plate, wash three times with PBS followed by three times with water, do this by adding PBS to 3 consecutive wells of a 6 well plate, and Ultra pure water to the last three wells of the plate, wash by dipping the coverslips into each well of the plate.

10. Dry

11. Add 1 drop of the embedding solution for fluoresnce microscopy on a leveled slide. The coverslip should be layed so that there are no air bubbles. Note that the cell side of the coverslip should be down, touching the embedding solution. With glue go around the edge of the coverslip to seal it. Then the glue is left to dry. This is light sensitive and should be kept in the dark immediately. Can be kept in the fridge for approximately 2 months wrapped in Aluminum foil.

VOLUMES OF SOLUTIONS REQUIREDFor 6 coverslips For 8 coverslips For 10 coverslips

Solution 1 50ml of PBS with Ca/Mg50mg 0.1% BSA25mg 0.05% Saponin

50ml of PBS with Ca/Mg50mg 0.1% BSA25mg 0.05% Saponin

50ml of PBS with Ca/Mg50mg 0.1% BSA25mg 0.05% Saponin

Solution 2 617.5µL Solution 132.5µL NGS

855µL Solution 145µL NGS

1,045µL Solution 155µL NGS

Solution 3 604.5µL Solution 132.5µL NGS13µL primary AB

837µL Solution 145µL NGS18µL primary AB

1,023µL Solution 155µL NGS22µL primary AB

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Solution 4 604.5µL Solution 132.5µL NGS13µL secondary AB

837µL Solution 145µL NGS18µL secondary AB

1,023µL Solution 155µL NGS22µL secondary AB

12.3. Actin Ring 12.4. Apoptosis12.5.

13.0 Other something14.0 Other something15.0 Other something

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