I am new to IHC and I am have not had success in staining with Bodipy 493/503 to get a clear image. I have attached an image to illustrate the problem that I am having. I have titrated the working amount from 20 μg/mL to 1 μg/mL to attempt to lessen the amount of “blotches”. However, this image is the lowest concentration that I have and it is still showing these “blotches”.
I am fixing the slides in 3% Formalin in PBS for 20 minutes at room temperature. Then I am washing 3 times with PBS with 5 minutes for each wash. Then I add my Bodipy for 30 minutes at room temperature. I wash this with PBS 3 times with 5 minutes for each wash with rinses before and after. Finally, I mount the sections with Vectashield with DAPI.
What is this interference that I am seeing and what could I do to improve this staining?
Hi, Zachary. First, BODIPY dyes are very hydrophobic. When put into an aqueous environment, they will quickly glob up and precipitate. The stock solution should be 1-10 mM in DMSO or ethanol. Upon dilution into aqueous media (at around 1-10 uM, and you are in that range; 1 ug/mL = 3.8 uM), shake the solution vigorously and then immediately apply to the sample.
I'm confused about your usage. BODIPY dyes are typically used for live cell membrane analysis, liposome imaging, or some other special lipid analysis usages. Using it on a fixed tissue section isn't typically performed.
Also, formalin (= methanol-stabilized 37% stock of formaldehyde) isn't typically used, since the methanol in it will delipidize the sample, thus reducing the ability of the BODIPY dye to label. Thus you would want instead to use a 16% methanol-free stock. Formaldehyde crosslinking, in general, will reduce the binding efficiency, since the binding efficiency requires a fluid membrane.
Once you have performed the label, you will want to wash more vigorously and longer.
If I may ask, what is the goal of using BODIPY? Given the limitations I mention above, there may be a better option for you.
Hi, Zachary. First, BODIPY dyes are very hydrophobic. When put into an aqueous environment, they will quickly glob up and precipitate. The stock solution should be 1-10 mM in DMSO or ethanol. Upon dilution into aqueous media (at around 1-10 uM, and you are in that range; 1 ug/mL = 3.8 uM), shake the solution vigorously and then immediately apply to the sample.
I'm confused about your usage. BODIPY dyes are typically used for live cell membrane analysis, liposome imaging, or some other special lipid analysis usages. Using it on a fixed tissue section isn't typically performed.
Also, formalin (= methanol-stabilized 37% stock of formaldehyde) isn't typically used, since the methanol in it will delipidize the sample, thus reducing the ability of the BODIPY dye to label. Thus you would want instead to use a 16% methanol-free stock. Formaldehyde crosslinking, in general, will reduce the binding efficiency, since the binding efficiency requires a fluid membrane.
Once you have performed the label, you will want to wash more vigorously and longer.
If I may ask, what is the goal of using BODIPY? Given the limitations I mention above, there may be a better option for you.
Hello, Jason. The goal of the staining with Bodipy is to visualize lipid droplets for their size and number. The literature that I have reviewed (https://doi.org/10.1371/journal.pone.0077774) has optimized the usage of this stain for lipid droplets. I used 10% neutral buffered Formalin, but to my knowledge this doesn't contain methanol, however I could be wrong. Other papers I have reviewed have used para-formaldehyde and formaldehyde with success. Additionally, Thermofisher technical support also stated that any aldehyde fixative should be appropriate for this stain. However, if these will affect the lipids, then I would need to change that.
So it seems that the most likely problem could be the aggregation of the dye before I stain the sections and then also the washing afterwards that could help clear some of these globs.
Hi Zachary,
I am also having the same problem of non-specific uptake of Bodipy dye by the cells. I was wondering if you were able to solve the same problem!
Nguyen Truong We don't know if the dye would be retained with fixation. Technically, it isn't fixable since there are no amine groups to be crosslinked by formaldehyde.
Article BODIPY 493/503 Staining of Neutral Lipid Droplets for Micros...
This protocol does BODIPY staining prior to fixation! Another member of my laboratory has also used BODIPY before fixation in the past.
EDIT July 2020: I've been following this protocol for months without issue now :)
Hi,
I also tried to stain lipid droplets with BODIPY after fixation with 4% paraformaldehyde in PBS and I had a similar result. Then I tried to stain before fixation (following the paper shared above by John Yn ) and problem solved.
https://bio-protocol.org/e1912
BODIPY 493/503 Staining of Neutral Lipid Droplets for Microscopy and Quantification by Flow Cytometry
This protocol maybe useful. However, the concentration of the BODIPY appears to be much lower than yours:
2 μM BODIPY staining solution:
- There is no background noise from the attached image
Hi all.
I am staining lipid droplets with bodipy and antibodies for ATPB and TSPO.
Following the advice of staining Bodipy prior to fixing, can I continue with blocking using 5%BSA+0.2 Triton-x followed by primary and secondary Abs?
Will the washing steps in between Abs wash away the Bodipy staining?
Thanks.
Hi John Yn. Can you tell me what is the DMSO concentration in which the BODIPY is diluted?
Best regards
Hi André Filipe Alves,
I used 90% DMSO and 10% PBS (I dissolved it first in PBS, my mistake...) for final concentration at 500µg/ml. I did the staining for lipid droplet with skeletal muscle at 25µg/ml for 1h is perfect, but for higher fat content (in my case ectopic fat) 1µg/ml for 1h was very good too (washing with PBST also good for that).
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