Histology and Pathology Method Forum

Beta-cell insulin immunohistochemistry problem! :(

Mon, 30 Jan 2012 17:59:24 +0000

Hi all,

Having a forum like this is really great and I am happy I could join - thanks to the team!

I have a question regarding immunohistochemistry for insulin and I am pretty desperate. I have the following protocol to stain insulin positive beta-cells for stereology, but somehow I can't get a strong signal (eg strong enough to take decent pictures of my slides and do some consistent counting). Info: The pancreas is collected, placed in 4%PFA overnight, then in 70% etOH for at least 24h and then embedded in paraffin.

~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Deparaffinization:
Xylene I, II, III, IV - each 6 mins
Ethanol 100% I, II, III - each 6 mins
Ethanol 90%, 70%, 50%, 30% - each 6 mins
1 minute of running tap water

Block
PBS-T with 5% serum from species in which secondary antibody was raised - 20 min at RT

1ary AB
2% serum block in PBS-T with 1:50 primary antibody (guinea pig anti insulin)

3 min wash in PBS-T

2ndary AB
2% serum block in PBS-T with 1:40 secondary antibody (rabbit polyclonal anti guinea pig IgG, with green fluorescence)

3 min wash in PBS-T

mount and fix with Vectashield+DAPI
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~

Ok so first I thought the problem was the embedding of the tissue itself, thus the many steps of deparrafinization and rehydration. This improved the overall signal strength. (I mainly did this because I noticed that when I do the whole protocol TWICE for a set of slides, then they look decent enough to count.)

Well so I don't want to repeat the whole protocol for each set of slides I do, as this costs time and reagents. Does anyone see an obvious flaw in the protocol?
Does anyone have the same problems with consistency in such protocols? I literally treat every slide the same way, process in the same amount of time, eg. I don't do too many slides at once, they don't dry out...and STILL it happens that slide a looks good but slide b shows nearly no staining at all...

I am really desperate as I have been trying to optimize this protocol for a while now,
please any help is welcome!

Many thanks,
best wishes,
Maya

Visualizing Biofilms

Fri, 06 Jan 2012 13:48:33 +0000

Hi,

Is there some way of staining biofilm cells, so that they can be directly visualized by eye and in a way that does not affect the viability of the cells?

thanks

Histology for mice lungs with metastases

Thu, 05 Jan 2012 17:22:55 +0000

Hello,

I am trying to do some histology studies for lungs that were extracted from mice. These lungs have metastases and I have followed the standard protocol used here for the dehydration, fixation and embedding. When cutting the sample using the microtome the paraffin was being correctly sliced, however we would get a hole on the area of the tissue. The tissue was not being sliced together with the paraffin at all. Does anyone have any idea why this was happening? Any help would be really appreciated.

The protocol used is shown here:

Histology Protocol

After extracting the lungs and the tumors from the animals, place them in a 10 % formalin solution (or 4% formaldehyde solution, diluted with PBS).
Store the tubes in the refrigerator overnight.
Replace the solution with 70 % ethanol. Store in the refrigerator. The specimens can be stored for several weeks.
In order to start the paraffin embedding place the samples in a beaker filled with 80 % ethanol for 30 minutes. The samples must be thin enough in order to fit the mold.
Transfer the samples into a 95 % ethanol solution and let incubate for 1h.
Replace the solution with a 100 % ethanol solution and incubate for 1h30. The beaker should be covered with parafilm in order to avoid any evaporation.
Incubate for 1h with Clear-Rite 3 (Richard-Allan Scientific) or xylene.
Place the samples in a beaker filled with paraffin. This beaker must be placed in a 60 ºC oven prior to adding the samples in order to assure that the paraffin is already melted.
Incubate for 1h30.
After incubation, pour a little bit of the paraffin solution into the bottom of the mold allowing to obtain a thin layer (approximately 2-3 mm thick).
Without waiting more than 30 seconds, place the sample in the mold paying particular attention to the orientation of the tissue. Place the orange lid on top of the mold and quickly fill it with the melted paraffin. Make sure there are no bubbles and that there is enough paraffin above the lid in order to leave approximately a layer of about 3-4 mm on top. After pouring the paraffin tap the mold against a surface a couple of times in order to make sure that the solution is uniformly distributed.
Place the molds on the cooling surface for 30 minutes.
Store the molds in the -20 ºC freezer overnight.
Remove the molds from the freezer.
Install the blade (which is stored in the wood box located in drawer below the desk where the microtome is) on the microtome.
Make sure the orientation is like 45 º for the blade. That can be adjusted in order to optimize the slicing of the specimens.
Adjust the cutting thickness to 7-8 microns.

c-fos protocol

Tue, 15 Nov 2011 06:16:03 +0000

I need c-fos protocol immunohistochemistry in non-free floatinf frozen sections, Any help??

Immunostaining of olfactory epithelia

Mon, 31 Oct 2011 09:47:42 +0000

Hey there,

Iam struggling with conditions of staining olfactory epithelia(Zebrafish). I did two attempts on fresh frozen tissue, dissecting in cold PBS then embedding in tissue tec followed by cryosectioning. When I perform the immunostaining I usually fix in cold Aceton for 15 Min, then wash the tissue and proceed to 1st AB without Antigen retrieval. My washing solutions have 0.1% Triton. When I mount the tissue and counterstain to observe it I always have kind of a mashy morphology. I also tried with Tween20 instead of Triton, still no change.

I know that fresh frozen tissue is more delicate and that nice morphology is hard to achieve. So what do you think about fixing in PFA for few minutes instead of Aceton? Will it help? I still want to ommit AR...Should I leave out the detergend at all?

Has anybody experience with epithelia or similar and can give few suggestions?

Thanks
Ivan

Histomount - Stuck!!!

Tue, 25 Oct 2011 09:43:29 +0000

We have been using histomount to mount stained sections..... however, the lid has become stuck shut! Does anyone know of any way to open it or is the whole bottle lost now?

Many thanks!

Forgot to mount slides after H&E, are they safe?

Mon, 24 Oct 2011 18:01:36 +0000

Hi, last Friday I did some H&E staining. Came back today (Monday) only to realize I forgot to mount them!

Stupid me... are they still usable...?

Yeah I feel pretty stupid right now...

abnormal epithelial cells in mouse mammary gland

Mon, 24 Oct 2011 14:48:48 +0000

Dear all,

I just begin to do mouse work. When I do H&E staining for paraffin embed section of mouse mammary gland, I found there are some abnormal shape epithelial cells, the nucleus are triangle or elongated not round (pls see the attached file, red arrow). Does anybody know what's these cells?

One thing I was thinking is that the mouse mammary gland undergoes involution, so the epithelial cells are "eaten" by surrounding nonprofessional phagocytes (they are also epithelial cells just because they have the ability to phagocytosis).

Thanks,

Bo

Attached thumbnail(s)

X-gal staining and IHC against b-gal

Sun, 25 Sep 2011 20:26:08 +0000

Hey guys,

I have a little bit of problem that I need your help to figure out what is going on.

I have this tissue in frozen block. this tissue is from a transgenic mouse with b-gal reporter.
(tissue was stained with X-gal before cryopreserved, so I can see there are blue cells in the tissue when I made section.)

I was using this tissue section and staining with some antibodies with DAB staining to see the identity of the blue cells.

Then, I just did staining of this tissue with anti-beta-galactosidase (DAB) antibody to make sure DAB staining and X-gal colocalize, but ended up realize some do, but some dont.

I mean that I can see some cells are blue (X-gal+) but DAB- (beta-gal negative), or DAB+ but not X-gal positive.

Does anyone see the similar results before, or know any publication reporting this? or any suggestive explanations for this?

BTW this tissue was isolated from animal,
Rinse with PBS,
Stained with X-gal solution,
Wash with PBS,
Fix with 4% PFA
and wash with PBS and proceeded to cryopreservation.

Any suggestions help, so please let me know

Thanks in advance

LDH measurement

Fri, 23 Sep 2011 02:42:00 +0000

How can I measure the LDH activity of a tissue performed as a biopsy?
And what can this data tell me about the level of necrosis??