Fix in 4% paraformaldehyde for 24 hours. Remove 4%PF and change to 1X Phosphate buffered saline (PBS). Change PBS once several hours later and again in a few days. Let me know if you need more detail.

You can used formol 4% with the same technique as above.

It depends on your proceeding. If you want to do immunohistochemical stainings you can use formalin fixation for most stainings. As mentioned above fix for 24-48 hours in 4% formalin. Thereafter you should embedd your tissue in paraffin without any further dilutions in PBS or alcohol.

If you are going to perform special stainings you should check first which preservation method suits best (Zinc fixation, cryo conservation ...)

you can use Bouin

department of Pathology : formol 10%

For usual istochemical stainings and immunohisatochemistry, you can fix tissues in 10% buffered formalin for 12-24 hours at room temperature. Then put tissues in 0.01M PBS.

Then you have to dehydrate them by changes in crescent ethanol concentrations, xylene and paraffin. Then paraffin embed them.

Only for H&E staining, the bouin's solution is enough, if wan to do do some special histochemical analysis then write down your target please

10 and 4 % formol is the same, as one is the abs. and the other is the vol. %age. Concentrated formol is 37%, so if you dilute it to 10%vol. it is 3,7 (~4)% abs..

PBS buffered is advisable, if you fix it for longer periods, which shouldn't be needed for the gut of a rabbit. Anyway to allow even fixation you should open the gut longitudinally .In order to prevent tough handling of the stiffer material after fixation, needle it to some cardbord, styro, or cork. You can also put it on cardboard (empty glove box) and it will stick to it, put it in the formol container ~500ml. And when you take it out the next day it will lie flat.

The PBS washing afterwards - I never tried it, and see not mandatory use in it.

Paraform and Bouin can be difficult if you want to do immunohistochemistry or extracts, very little pieces in Paraform are needed in case you'd like to do electron microcopy.....

for H&E saline 10% buffered formaline will excellent results.

To prepare it :

Add one volume of concentrated formalin to nine volumes of normal saline(0.9%).

Remember that intestinal mucosa is very fragile and quickly autolyzes.So

immediate fixation after sacrifice of the animal is essential.You have to put

some formol saline inside the lumen of the intestine and then put it in the formol saline

at least 5 times its volume. Leave it for 2-3 days.

10% buffered formaldehyde should work well for most tissues. Cut your blocks to be about 2 - 3 mm thick at the most so that they are well permeated by the formaldehyde (10% is the same as 4% because the full strength of formaldehyde is 40% so that when most of us talk about 10% it is in fact 4%). Fixation for 24 hr should be ample time after which you should be able to process your samples. Depending on the size of the material you may have to experiment with an overnight run or a short run if the overnight processing is makig the tissue too brittle and dificult to produce good sections

accord to method advised by prof Alam Nafadi head of electron microscopy unit assuit university - fast animal 24 h then use cardiac perfusion with saline followed by 10% neutral buffered formalin - tie the oseophagus , and rectum fill the whole gut with 10% neutral buffered formalin - imerse the whole tied gut in wide container filled with the same fixative - leave 2-3 hours then open and cut either tranvesres or longtudinal accord to the obvjective wash througly in saline to remove the content =then fix again for 24 hours before processing

for EM wash the tube with saline - tie the part u need from the 2 ends - then fill with gluteraldhyde in the same way ( best regard)

Bouin's fluid is very good. It is a good fixative and preservative. You can get excellent results.

I made best preparation with neutral 10% formalin. after that staining is very easy and benefit for microscopical evaluation.

Acccording to me, formol 4 or 5 % is the best fixation.

May be the best option is that offered by Soad Ali from KAU

If it is only H&E you are doing, no other follow up, it really isn't overly important, H&E is not that picky. I tend to use 4% formalin in PBS, it is what we have on hand in quantity. Since intestine is not very thick you shouldnt need to worry about penetration, it might be a good idea to squirt some down the lumen. Use plenty of volume and allow 24 hours. If you are doing immuno, see what the manufacturer recommends.

I would suggest the use of 10% (final is 4%) buffered neutral formalin. Please give one change of formalin after 4-6 hrs and 24 hrs after initial fixation, proceed with paraffin-embedding. This is for H&E. However, for doing special stains (immuo or histochemical) please check with the manufacturer's instructions.

Bouin's fixative will be very good for your work (24hrs).

10% buffered fomoline is the best. You can use the formaline fixed material for H&E staining, special histochemical stains, most immunohistochemistry stains.

I will also suggest that, you go for the Bouin's fixative. As your doing nutritional analysis it is very good fixative for it. At least for H & E staining it is good and if you want to go for protein (bromo-phenol blue staining, Ninhydrin schiff's staining), carbohydrate (PAS staining) or lipid (Sudan black B/ oil red O staining) analysis/staining it is fairly good. Fixation time 24 hrs. but while dissecting give cut to the intestine and open it so that intestinal villi can be completely exposed to the fixative and it will also help you during the wax infiltration and impregnation. Section thickness upto 5-7 micron will also gives better discrimination of cell type.

I would recommand 10% buffer formalin for paraffin embedding which allow you for further investigation (immunohistochemistry) without any crossreaction.

If you need to preserve cytological details I would suggest to fix in glutaraldehyde, but you need to dice the sample as small as possible for better preservation of cells.

10%buffered formaline

It depends: if you want only to see the best morphology, formalin is the choice. If you want to perform different studies (melcular biology for example) you can use an ethanol based fixative. A good compromise should be the "cold"fixation in formalin at 4°C (24 hours) where you can observe a good morphology together with a good DNA and RNA extraction.

it actually depend on what you are going to study. if you are going to study enzyme histochemistry or immuno-histochemistry or immuno histology study you can use microwave fixation [heat fixation]. if you do simple norphology studies it is bouin fixative and if you go for electron microscopy it is glutardehyde - aldehide fixative. use of formaldehide as 10% buffered Formol saline or formol saline will destroy antigens and enzymes and need antigen retrieval. use of 10% formol saline will also destroy the mucin, glycogen, fat etc

10%buffered formalin is very good & available fixative for histologic studies

Open the bowel and put it in 10% buffered neutral formalin for paraffin embedded tissue.

i have already suggested to use Bouin's fixative than 10% formaldehyde for best results . please try

I prefer to use 5% buffered formol saline and change it twice after 24 hours. but gut pieces should be gently flushed. Bouin's does well but should be replaced with 70% alcohol after 4 hours.

Bouins fluid or 10% formalin. However, be sure to use do it in the approximate ratio of 1 part of tissue to 9 parts of formalin for best results

fix the tissue in either 10% neutral formalin or aqueous Bouins fluid for 24 hrs. the ratio of tissue and fixative should be 1:10. after thorough washing in running water for 24 hrs. dehydrate the tissue in graded alcohol series.After dehydration it is advisable to clean the tissue in benzene instead of xylene and then proceed for embedding in paraffin.

the best fixation for the tissue is 10% neutral formalin for 12-24 hrs. then wash the tissuel in runing tab water for 12 ,hrs. dehydreat the tissue in graded alcohol series ( 70% - 80%- 90% - 95%- 100%)..After that clear the tissue in benzene instead of xylene and then proceed for embedding in paraffin.

y Con't u try in4% para formaldehyde

Open bowel, not stretch it and fix it rapidly in buffered neutral 10 % formalin.

the fixative most widely used is the neutral buffered formalin to 10%, for a time that must vary depending on the type of reaction must be submitted to the histological section. For histochemical fixed for 1 week or more, followed by a washing in tap water for 1 hour, being the reversible fixation in formalin a washing may compromise the morphology, then continued as per routine.

Buffered formalin 10% is excellent.

For histology, you can fix in Bouin 's fixative whereas for histochemical you can go for PLP.

Please look up the paper titled " What is the Best Procedure to Remove

Formalin Pigment from Formaldehyde-Acetic

Acid-Alcohol Fixed Tissues? " with the following abstract:

There has been some disagreement as to whether formalin

pigment is bleached by potassium permanganate/oxalic

acid (PPO). This study analyzed the bleaching characteristics

of PPO on tissues fixed in formaldehyde-acetic acidalcohol

and compared PPO with other bleaching agents and

conventional formalin pigment removal methods. It was

found that PPO removed formalin pigment. Other bleaching

methods (hydrogen peroxide, periodic and chromic acids)

also removed the pigment to some extent. Garvey’s pigment

removal procedure was more effective than alcoholic picric

acid for the routine removal of formaldehyde-acetic acidalcohol

associated formalin pigment. ( The J Histotechnol

33(3):109–111, 2010)

Sir

You can use FAA as the best fixative for the purpose. Because I used FAA for the histological studies in my lab.

We had excellent results for H&E stained paraffin embedded sections of the dog and rabbit intestines with modified Bouin's fixative, The preservation and staining (colours of both acidophilic and basophilic sunstance) was so good that the sections prepared in 1970, are still used for teaching veterinary histology.

Gastro-intestinal tracts are very difficult to fix because of the presence of bacteria and digestive enzymes.

Formalin is an aqueous solution of formaldehyde and concentrated formalin is about 40% formaldehyde. 10% formalin (about 4 % formaldehyde) is a good fixative but process of fixation takes time whereas Buin's fixative fixes rapidly but penetration is not as good as formalin. We experimented to mix formalin with Buin's and got excellent results. Buin's fixative formula is : Saturated aqueous picric acid 75 ml, 40% formaldehyde (concenrated formalin) 25 ml and glacial acetic acid 5 ml. 10% formalin is 90ml of water and 10ml of concentrated formalin. Mix the same volume of the two fixatives. The 10% formalin is used in neutral pH using the buffer but for this mixture, there is no need to adjust the pH.

You can use 10% formalin to fix soft tissues . But for vertebrate tissues only bouin's fixative is used to get good histological sections after embedding in paraffin wax and then dehydrating with xylene and alcohol series and staining with haemotoxylin and eosin for better results of histological sections .

Hi Yasser

I advise you to do the following steps:

1. After dissection, cut the digestive tube into 4 parts (duodenum, jejunum, ileum and colon)

2. Wash with saline (0.9% NaCl)

3. Re-cut each of the mentioned parts again into many small pieces (2-3 mm) and wash again with saline

NB: If you want cross sections, leave the parts as small tubes, but if you want longitudinal sections you should open the small pieces and lay them down in the mold.

4. Fix the tissues in buffered neutral 10 % formalin (to avoid acidity of formalin) for 48 hours.

NB: don't forget to put tissues in different containers labelled duodenum, jejunum, ileum and colon.

5. Wash tissues (put in gause) in running water for 4 hours to remove any traces of formaline.

6. Dehydrate in ascending grades of alcohol (50-70-95-100)%

NB: 2 changes of absolute alcohol and time dependes on the tissue thickness

7- Clear in xyelene (two changes)

8- Infilterate in wax-xylene, wax1, and wax2

9- Make your paraffin block

Good luck

I think now you have excellent options , you may like to use that of Gabir Ramadan.It depends on what you like to do with this tissue.

I will select Bouin"s fluid for almost every routine histological work, exception is the kidney tissue. I also would like to say that oesophagus and stomach are parts of the digestive tube (Gaber Ramadan)