It seems to be quite away from the nucleus and indeed too big and irregular to be within the endosomal compartment. I agree with Patrice, it could be protein agregates possibly within lysosomes. I would try to overexpress chaperones and/or reducing plasmid amounts.

Claire

By the way,the spots labeled in red are formed by my target protein,and the blue ones are labeled marker.

Thank you,I will try another endosome marker next time. Do you mean exogeneous protein expressed in mammalian cells will sometimes aggregate like inclusion body in E.coli?

Indeed,I am trying to localize a protein in host cell,by overexpress it with a myc-tag,as we can not find a proper primary antibody to detect the corresponding endogeneous host protein.However,after IF,the staining is quite weak,some of them can't even be discerned from the background,but others are the case shown in the above image attached.As I vortex the DNA/Turbofect mixture,and rock the dish gently after transferring the mixture to the cell culture to assure the even distribution of plasmid,so I am puzzled by the outcome of IF.What should be a successful transfectant looks like?Take EEA1(early endosomal antigen 1)as an example,should most of the transfectant show colocalization between Myc-EEA1 and other early endosomal marker?

Just to add something, with respect to inclusions, you do get them in human cell lines. I have seen them on many occassions, also I am very wary of overexpression as you are biased to the locality of the protein. Why do you require to overexpression? Why not look at the endogenous protein! Is it access to the actual cell type in which your protein is present?

Cheers gareth

To：Gareth Sullivan

Because we can find proper primary antibody for endogeneous protein,so we chose to add a Myc-tag at it's N-terminus.And I believe HeLa cell will express my target protein.Thank you a lot.

The proposal of the endosomal localization it is not so clear for me, as the endosomes are formed mainly containing extracellular proteins or membrane proteins after a stimulus. Its more probable that your protein, as overexpresed, are located in vesicles from RE or Golgi. The ER has mechanisms to discard proteins misfolded (due for overexpresion) and this could one of them. Have you look for co-localization in this organelles? For ER you can use an anti-Calreticulin or anti-Calnexin antibodies, for example. For Golgi there are many markers you can use.

I thinks these are not vesicles...too big, I would suggest these are proteins aggregates.

have a look the extend of aggregation you might have with GFP tagged protein. two cells with large aggregate (other cells are Microtubules bundles by the PRC1 proteins

I am not sure that there are endosomes. To make picture more clear, you should make co-staining with the other markers, at least nucleus (DAPI) may partially help. But the best way to clarify results is co-staining with organelle markers like endosomes (http://www.cellsignal.com/products/2143.html) or endoplasimic reticulum. Good luck!

Thank you all so much.

Also, to me, those puntae don't look like endosomal or vesicular inclusions. I'm working on aggregating and misfolded proteins (cytosolic, as well as lysosomal aggregates) and we use overexpression as one tool to trigger intra-cellular aggregation.

One additional way to gain information, if the co-staining for organell markers is not painting a clear picture, is to express the protein fused to GFP and observe the stability of the punctae upon various concentrations of detergent. If they are just held together by a vesicular membrane, even mild detergents (0.75% NP-40 or 0.5% TritonX-100) should dissolve all the fluorescent punctae. In such an approach with non-fixated, fluorescent proteins, you might also try to photo-bleach a part of a big fluorescent blob and observe the regeneration of the fluorescence - called Fluorescence Recovery After Photo-bleaching (FRAP). The longer this regeneration takes, the less mobile the fluorescent molecules are.

Moreover: If you have to over-express your protein to study it within the cell, you also might want to use the lowest expression-rate suitable for your needs. Hence, try other, weaker promoters or establish a stable (virally transfected) cell line with a subclone selected for weak and soluble expression of your protein (due to just one insertion event of the recombinant gene).

Also, a GB1-tag fused to the protein often increases its solubility. However, whatever tag you use, always think of the proper controls!

Good luck!

I wonder which marker you've used to stain endosomes. They look a little too big for endosomes, they resemble more the lysosomes. I also wonder what kind of a protein you've expressed. Some Rab GTPases have the property to change the size of endocytic vesicles when they are overexpressed since they mess up membrane fission/fusion events.

Hi,

I have been doing some experiment using co-localization of protein by multicolor Immunofluorescence (mIF). It's clear looking at your picture is present a co-localization of two different color (Blue and Red). Indeed, the violet color represent the merge between Blue and Red. However, should be necessary to understand whether we are looking for a effectiveness co-localization or there are some pitfalls in your procedure.

At first impression seem to be co-localization.

Al the best

Niccola

to：Philipp Ottis

I totally agree with you on that lowest expression-rate plasmids would be better when analyzing the subcellular localization of my target protein.However,it seems difficult to detect low abundance protein with antibody,at least in our lab,we fail to do that in many cases.

However, I don't agree with you on that- " If they are just held together by a vesicular membrane, even mild detergents (0.75% NP-40 or 0.5% TritonX-100) should dissolve all the fluorescent punctae."Indeed I permeablize the cells with 0.5% Triton X-100 for 3 rounds,and 10 mins for each round.If 0.5% Triton X-100 will dissolve the membrane of endosome, then the membrane marker(stained blue) ought to permeate throughout the cell plasm,or wash away subsequently.

Anyway thank you very much.

Could the inclusions be mitochondria? The size would fit. There are mitochondria-specific markers to test this possibility.

Dear friend,

Could this be lysosomes... You may try Lysostracker red from Molecular probes...

I do not feel they are related to the mitochondrial compartment.

Patrice

It seems to be quite away from the nucleus and indeed too big and irregular to be within the endosomal compartment. I agree with Patrice, it could be protein agregates possibly within lysosomes. I would try to overexpress chaperones and/or reducing plasmid amounts.

Claire

Dear Arrhenius,

was the image obtained from a confocal microscope? If it is not, have a look on your cells in a confocal microscope; if you see a more ring-like structure, the stained structures may represent lipid droplets.

Best Stefan

What controls have you done and what did they look like? I've never worked with HeLa cells, what two things are the two colors showing? To what organelle do you expect the protein to localize? You could try targeting your protein to a different organelle, then if the puncta move, that would be consistent with you having found the original organelle, while if they don't that would be consistent with aggregation.

Before moving on to more advanced markers, a better handle on the nature and structure of the inclusions would help. They are definitely not fibrillar cytoskeleton, despite distribution right out into the filopodia. Intensity and shape suggest they are membrane-bound or at least have a hydrophobic surface; that they are variable in size and peripheral suggests either endocytosis, exocytosis or a relationship to filopodia metabolism. Look at the preparation unstained and note any autofluorescence, maybe with diffraction filters; I strongly doubt mitochondria, but you can do an easy stain with para-phenylenediamine to check.

You might try Nile Red vital staining for lysosomal phospholipid inclusions, strongly fluorescent in a hydrophobic environment (Greespan et al., J Cell Biology 100: 965). See Hogue et al. (J. VIrol. 85: 9749) who describe lipid raft coalescence in virus-infected HeLa cells.

More on point, monodansylpentane (MDH) is a high contrast blue-fluorescent marker used to track lipid droplet trafficking in live cells (Yang et al. PLoS ONE 7(3): e32693). The lipid association is suggested by shape and refractility of your inclusions. A fabulous stain to show lipid conformation is osmium tetroxide, use in most structural electron microscopy, but can also be used for light microscopy: at the EM level, pure lipids can be judged by level of saturation (intensity of osmium staining) and whether the inclusion is lamellar or not (membranes). A lysosome looks very different from a lipid deposit.

Why not lyse (distilled water) your attached cells, and either filter the various sized debris or just centrifuge at low speed through a gel to create layers? It would be so much easier to characterize the bodies if isolated. If lipids: they’ll float! A protein granule will spin down like a rock.

It is really difficult to offer an interpretation of these images without more information. What is the organelle for which you are staining? With the information provided, I agree with previous comments, it looks lysosomal. Are you staining the organelle marker with an antibody? If so, it is possible (and likely) that the antibody staining has some non-specific labeling in addition to labeling the organelle. Hence, I would not be concerned about some level of non-overlap of the two markers, the two markers seem to have too much overlap to be a coincidence. I would try a few other ways to label your organelle. For example, if you are indeed looking at lysosomes, LysoTracker is a good choice, as suggested above. I would not bother with trying to biochemically isolate these structures, as this would require a lot more work than simply doing some additional fluorescence staining conditions and controls, without being any more informative.

One might consider increasing the resolution a bit and do electron microscopy in combination with immuno labeling.

Transient overexpression often overloads the cell and can result in aggregates or induction of the unfolded protein response. If your protein has a signal peptide and is trafficked through the ER it is likely to overload the capacity of the ER to handle it. You could try titrating the amount of plasmid transfected. Even for large plates (100 mm) we sometimes us as little as 100 ng of expression plasmid so as not to overload the cells.

hi there,

Have you ever tried to co-localize those cytoplasmic spots with stress granules-related proteins, such as TIA, TIAR or FUS?

Some reagents that we use for transfection can cause stress on cells.

For instance, lipofectamine2000...

I think it's worthy to check it, just in case.

Best wishes,

@lex