fixation of alginate beads - formaldehyde and CaCl2 --> reaction? (Jul/02/2007 )
I want to fix my cells in alginate beads. It is necessary to add CaCl2 to prevent my beads from dissolving, but can I just add some CaCl2 to 4% formaldehyde or will I then trigger some reaction (I'm not really a chemist).
Ca++ bridges COOH groups on alginate to cross-link the polymer. You should have Ca++ in the system already when you made the beads with cells entrapped within. Formaldehyde does not react/interfere this process.
Aspergillie,
My beads fell apart when stored in normal formalin for about 2 days, so I added some CaCl2, which helped. I think it diffuses out with time, as it equilibrates with the fixative solution. I've read some folks use Barium instead of Calcium to form beads used for histology, which is supposed to prevent them from dissolving. But now I'm using a different fixative, and am quite happy with it. Here's the recipe for 15 ml from my protocol book, that I use for chondrocytes in alginate beads... it's a complicated, but works really well for both cellular and ECM preservation for light and EM images. Also if embedding in paraffin or OCT, this makes the beads dark-brown in color, so they're easy to see in the block when cutting.
Materials
- 2% (v/v final) 32% paraformaldehyde EM grade from EMS #15710 =0.93ml
- 2% (v/v final) 50% glutaraldehyde EM grade from Sigma #G7651 = 0.6ml
- 0.7% RHT (w/v) final RHT=Ruthenium (III) hexamine trichloride from Alfa-Aesar #10511 - !!!Add fresh immediately before fixing!!! 0.105g
- 2% Sucrose (w/v final) MB grade Sigma# S0389 = 0.3g
- 0.1M final Sodium cacodylate trihydrate Sigma #C0250 =7.5ml 0.2M (2x) stock
- 10mM final Calcium Chloride MB grade- Sigma # C3306 use from 102mM Gelling buffer = 1.5ml
- ddH2O = 3.6ml
!!Wear Gloves!! - this stuff is toxic - Aldehydes, Arsenic and heavy metals Oh my!
Procedure
1) Prepare a 2x stock of buffer: cacodylate (0.2M), 20mmM CaCl2, 4% sucrose in ddH2O; pH to 7.4 then....
Vortex til dissolved; store at 4`C until use.
Day of use (~4 hours before use)
2) Add paraformaldehyde in fume hood-->vortex briefly
3) Add glutaraldehyde in fume hood--> pipette slowly as 50% Glutaraldehyde is viscous, and pulse ~20 times with pipette to mix
4) Add ddH2O to final volume
5) hold in dark, on ice until ready to use
6) weigh out required amount of RHT --> transfer to a 1.5 ml eppendorf and wrap in foil
7) immmediately before adding fixative to beads, add RHT, mix to dissolve. Initially solution will be yellow, but changes to a dark brown in a few minutes...this is normal.
8) Wash beads with 10mM CaCl2 buffer-->add 10 volumes fixative per volume washed beads eg 1 bead = 40ul, so 400ul fixative
9) allow to fix for 2 hours at 4`C, then wash beads twice in 1x Cacodylate/CaCl2/Sucrose. Hold in this buffer overnight at 4`C and then change to fresh buffer; beads can be stored @ 4`C for ~2 weeks.
Hope this works for you, aspergillie. Cheers!-JAH
My beads fell apart when stored in normal formalin for about 2 days, so I added some CaCl2, which helped. I think it diffuses out with time, as it equilibrates with the fixative solution. I've read some folks use Barium instead of Calcium to form beads used for histology, which is supposed to prevent them from dissolving. But now I'm using a different fixative, and am quite happy with it. Here's the recipe for 15 ml from my protocol book, that I use for chondrocytes in alginate beads... it's a complicated, but works really well for both cellular and ECM preservation for light and EM images. Also if embedding in paraffin or OCT, this makes the beads dark-brown in color, so they're easy to see in the block when cutting.
Materials
- 2% (v/v final) 32% paraformaldehyde EM grade from EMS #15710 =0.93ml
- 2% (v/v final) 50% glutaraldehyde EM grade from Sigma #G7651 = 0.6ml
- 0.7% RHT (w/v) final RHT=Ruthenium (III) hexamine trichloride from Alfa-Aesar #10511 - !!!Add fresh immediately before fixing!!! 0.105g
- 2% Sucrose (w/v final) MB grade Sigma# S0389 = 0.3g
- 0.1M final Sodium cacodylate trihydrate Sigma #C0250 =7.5ml 0.2M (2x) stock
- 10mM final Calcium Chloride MB grade- Sigma # C3306 use from 102mM Gelling buffer = 1.5ml
- ddH2O = 3.6ml
!!Wear Gloves!! - this stuff is toxic - Aldehydes, Arsenic and heavy metals Oh my!
Procedure
1) Prepare a 2x stock of buffer: cacodylate (0.2M), 20mmM CaCl2, 4% sucrose in ddH2O; pH to 7.4 then....
Vortex til dissolved; store at 4`C until use.
Day of use (~4 hours before use)
2) Add paraformaldehyde in fume hood-->vortex briefly
3) Add glutaraldehyde in fume hood--> pipette slowly as 50% Glutaraldehyde is viscous, and pulse ~20 times with pipette to mix
4) Add ddH2O to final volume
5) hold in dark, on ice until ready to use
6) weigh out required amount of RHT --> transfer to a 1.5 ml eppendorf and wrap in foil
7) immmediately before adding fixative to beads, add RHT, mix to dissolve. Initially solution will be yellow, but changes to a dark brown in a few minutes...this is normal.
8) Wash beads with 10mM CaCl2 buffer-->add 10 volumes fixative per volume washed beads eg 1 bead = 40ul, so 400ul fixative
9) allow to fix for 2 hours at 4`C, then wash beads twice in 1x Cacodylate/CaCl2/Sucrose. Hold in this buffer overnight at 4`C and then change to fresh buffer; beads can be stored @ 4`C for ~2 weeks.
Hope this works for you, aspergillie. Cheers!-JAH
Thanks a lot!