Dear all: It is interesting this Topic "Histology" was opened just a few day ago. therefore either are not questioned often since otherwise the topic had to be opened years ago. I guess that the professionals out there don't like to discuss about. Other issues dealing with "histology" have been questioned and replied to in other ResearchGate Topics like , and previously. Now following my personal reply to So-Jin KIM (just focusing on prep. issues like : Dear So-Jin, guessing "protocaking blocks" (see your question) was misspelled or rapidly written should be "protocol books" the answer is: Yes there are a lot of books (even "Handbooks of") on . I am not quite sure about but assuming your being a “novice” to Histology or Histotechnology”. If I am wrong with this, apologize. There are some questions to ask in first place (and perhaps we’ll get some answers?): 1) do you need your / only H&E-sections for SCORING = evaluating the MORPHOLOGY only and reporting / documenting tissue damage of the organs heart, lungs, kidneys, and liver, OR 2) do you need further (specific) stains like (classical Enzyme-) Histochemistry-Immunohistochemistry-specimens for molecular studies, electron microscopy, ...etc?. As some colleagues in their replies stated above, 10% (phosphate) buffered FORMALIN (note: WHAT IS THIS REALLY?? see following text) usually is a common used fixative to get started for most of the tasks listed above (but NOT usable as a primary fixative for TEM-studies). NOTE: 10% "Formalin" (which is a trivial trade name of the company ) = ca. 3,7-3,8% hydrous formaldehyde solution [formaldehyde is a GAS solvable in water up to a maximum of 37-38% end concentration, so “100% Formalin” = 37-38% hydrous !formaldehyde solution!]. This fixative is a so called “allround" fixative, developed long ago by empiric experience and functions as good in the “Death Valley” as in “Kathmandu, Nepal”. It is also of essence you might optimize your fixation protocol before starting your work (one can find not only in books but also searching by Google….) for several kinds of LM-high resolution morphological studies (so you would be advised to use Na-Na-PO4-buffered formaldehyde solution (instead of only using of PBS…> as the buffer, freshly prepared formaldehyde solution made from PFA [=para-formaldehyde powder), etc.,etc.]. The scale of producing is open here…. BLOCK SIZE of tissue should not exceed say 2 (W,L) x 3 (W,L) x 0.5 (THICKNESS) cm (yes: centimeters), due to the fact that formaldehyde solution penetrates at a low rate into the tissue (in the cold = 4 degr.C at least 5-10 times slower than at room temperature, but 3-5x faster than other aldehydes, like glutaraldehyde for instance). Therefore, ONE dimension of the tissue specimen / tissue slice (slab) always should not exceed 0.5-1.0 cm for optimal fixation of the whole! Usually, pathological material (i.e. surgery specimens) is fixed at room temperature, sometimes – to speed up – also at slightly elevated temperatures (up to 37-40 degr. C, 2-3 hrs) and sometimes also combined elevated temp. & under pressure (automatic devices). For evaluating tissue damage you should know first how to approach, and esp. choosing a sectioning plane through the organ you study which will show the matter you are after in a correct manner (anatomical issue). In the case of human heart or human kidney or human lung it is worth to know also that section dimension could be as large as 5-10 cm in diameter (and more): so you have to choose again which area you would like to evaluate or at least how you’ll divide the gross portions of your organs correctly (but if it is mice – what we don’t know for now – or rat or, or…? this might be no problem at all). FIXATION TIME (what is too short, what is o. k, what is too long?) in the range of at least 6 hrs RT (room temperature, use of a specimen probe rotator device) for a block say 10:10:5 mm I.M.H.O. as a minimum perhaps is ok ; immerging the spec. block too short produces a block unfixed in the center portion and therefore you will not be able to get a section of good quality. I agree with Wayne Briner in using always fixative 20-25 x the volume of the specimen. Tissue rotated during fixation in a specimen rotator device will be fixed easier and more thoroughly than tissue which just lies at the bottom of a phiole / sample glass tube . STORAGE in fixation solution MIGHT be no problem for ONLY morphological evaluation of tissue damage, but there are some CAVEATS: formaldehyde solution has been reported to act as dehydrating agent, if you store them too long in the fixative your specimens could get too hard and therefore – when embedded afterwards e.g. in Paraffin – the fixed tissue gets brittle properties which makes sectioning difficult, especially also if you use too long dehydration steps (We have not heard/read about dehydration or embedding issues (alcohols, or acetone, Paraffin or cryo-techniques so it would be interesting to get knowledge about your thoughts). Formaldehyde in slightly alkaline conditions (= pH 7,2 to 7.4; and not only those) is known to polymerize slowly but consistently all the while… does harden the specimen block…. so it might be advisable to lengthen the washing steps (with an appropriate buffer solution) after fixation. Also, formaldehyde solution left with the tissue produces formic acid (one of some byproducts) which is neutralized by the used (thats why we have to buffer the FA-solution, otherwise you run into acid fixation which distorts your tissue). It is right to say that also ARCHIVAL SPECIMENS stored for years in buffered Formaldehyde solution have been processed into paraffin blocks successfully, but this might not be the rule or be useful either. In general storage up to one week (or 2, especially at 4 degrees C) is acceptable and has been reported often as to be not damaging tissue per se. SPECIFIC FIXATIVES for different organs: YES, but usually this is an issue if you do vascular perfusion of whole organs. Then it might be advisable to adjust the fixative (also with additional cationic substances like CaCl2, MgCl2, etc) for perfusing brain, another for kidney, and another one for liver…. Note: Storage of tissue after fixation in 0.01 M PBS, as suggested by Lorena Salvatore I would not recommend due to hypo-osmolal(r)ity (unless the used PBS here is 10x concentrated): the usual concentration of storage buffer I.M.H.O.(in my honest opinion) is 0.1 to 0.15 M (0.13 M PO4 = equals to 300 mOsmol = the osmolarity of blood). Concerning "...1 day before mue.." in your initial question: I don't know what you mean with this but it could be "before dehydration, before …??). I know that my lengthy note perhaps is not very welcome (I don’t want to proselyte someone here) If you have more specific answers or also specific questions, please don’t hesitate to send an e-mail to [email protected] [EDITED 2016-08-02: DELETE: [email protected]   since meanwhile retired:  SEND YOUR REQUEST if needed to [email protected], thank you!] or to post this in ResearchGate-Topic . Best wishes and good luck with your work, Wolfgang MUSS, PhD, EM-Lab of Univ. Inst. Pathology, SALZBURG, Austria-EUROPE

NB: [only e-mail address edited, see in text above:

EDITED 2016-08-02: DELETE: [email protected] since meanwhile retired: SEND YOUR REQUEST if needed to [email protected], thank you!] 

First, what is your sample source? For human tissues: fixed 10% formaldehyde 4~ 12 hrs, room temperature (depend on your tissue size, your tissues can pack into a cassette that is the often standard which used in the clinical or research.) For mouse tissue: If your tissues need to do more study, for example, the IF Stains. I suggest you fixed your tissues in the 10% paraformaldehyde overnight -> washing PBS Buffer 3 times, 5 min -> 10 % sucrose solution overnight -> one half pack into OCT cryo-gel, stores in -80 degree and the other half made become paraffin block.

the fresh tissue is preparing for IF stains or IHC. The paraffin tissues is staining H&E stains or any special stains or IHC...

H&E is not that picky. 10% phosphate buffered formalin, pH about 7.4, room temp. If you let it fix more than a day the tissue handles easier. Longer is not a problem either. Have done histology on tissue that has been archived for years. Couple of things to keep in mind. Make sure you have plenty of fixative in your sample container. The rule is 30 times the volume of the specimen. Also formalin only penetrates a few mm, make sure your sample is no more than 4-5mm wide or the center will not fix properly. If you need an entire large organ fixed (ex brain) then you must use vascular perfusion.

I have done histopatholy of muscle tissue using only frozen sections

I agree with Wayne Briner: 10% formalin fixed tissues will be OK, but if you will do also immunohistoxhemistry, it's better to fix tissues for 12-24 hours maximum. Then put them in PBS 0.01M. Tissues can be stored for 2-3 months at room temperature.

Best results we can get with aqueous Bouins fluid fixative for 24 hrs, then washing with water to remove extra fixative.

you may use 4% paraformaledhyde made in 0.01 M phosphate buffer for 24 hours at 4C, but just be sure the Ph OF THE FIXATIVE IS ABOUT 7.4

Remove and Fix the tissue immediately in10% buffered formalin at room temperature for 24 hours which is ok for most of the tissues. Further it is the washing and dehydrating that is important and clearing and making of the blocks and cutting of the sections using a microtome. The sections are then stained with H AnD E after dissolving the paraffin and hydrating the sections . In fact each step is critical and needs to be done meticulouly. Anatomical changes thus can be recorded . In addition scoring can be done by using histochemistry methods also which should reflect functional morphology.

You also need to worry about the thickness of the fabric. To fixation for 24 hours is required that the thickness is less than 3 mm

For fixing, sample’s size should be about 1cm2, and then fixed in 10% formalin buffered. For using in the lung, place the sample in vacuo, with formalin, until there is no air inside the lung. After this, use histologic routin.

Dear all: It is interesting this Topic "Histology" was opened just a few day ago. therefore either are not questioned often since otherwise the topic had to be opened years ago. I guess that the professionals out there don't like to discuss about. Other issues dealing with "histology" have been questioned and replied to in other ResearchGate Topics like , and previously. Now following my personal reply to So-Jin KIM (just focusing on prep. issues like : Dear So-Jin, guessing "protocaking blocks" (see your question) was misspelled or rapidly written should be "protocol books" the answer is: Yes there are a lot of books (even "Handbooks of") on . I am not quite sure about but assuming your being a “novice” to Histology or Histotechnology”. If I am wrong with this, apologize. There are some questions to ask in first place (and perhaps we’ll get some answers?): 1) do you need your / only H&E-sections for SCORING = evaluating the MORPHOLOGY only and reporting / documenting tissue damage of the organs heart, lungs, kidneys, and liver, OR 2) do you need further (specific) stains like (classical Enzyme-) Histochemistry-Immunohistochemistry-specimens for molecular studies, electron microscopy, ...etc?. As some colleagues in their replies stated above, 10% (phosphate) buffered FORMALIN (note: WHAT IS THIS REALLY?? see following text) usually is a common used fixative to get started for most of the tasks listed above (but NOT usable as a primary fixative for TEM-studies). NOTE: 10% "Formalin" (which is a trivial trade name of the company ) = ca. 3,7-3,8% hydrous formaldehyde solution [formaldehyde is a GAS solvable in water up to a maximum of 37-38% end concentration, so “100% Formalin” = 37-38% hydrous !formaldehyde solution!]. This fixative is a so called “allround" fixative, developed long ago by empiric experience and functions as good in the “Death Valley” as in “Kathmandu, Nepal”. It is also of essence you might optimize your fixation protocol before starting your work (one can find not only in books but also searching by Google….) for several kinds of LM-high resolution morphological studies (so you would be advised to use Na-Na-PO4-buffered formaldehyde solution (instead of only using of PBS…> as the buffer, freshly prepared formaldehyde solution made from PFA [=para-formaldehyde powder), etc.,etc.]. The scale of producing is open here…. BLOCK SIZE of tissue should not exceed say 2 (W,L) x 3 (W,L) x 0.5 (THICKNESS) cm (yes: centimeters), due to the fact that formaldehyde solution penetrates at a low rate into the tissue (in the cold = 4 degr.C at least 5-10 times slower than at room temperature, but 3-5x faster than other aldehydes, like glutaraldehyde for instance). Therefore, ONE dimension of the tissue specimen / tissue slice (slab) always should not exceed 0.5-1.0 cm for optimal fixation of the whole! Usually, pathological material (i.e. surgery specimens) is fixed at room temperature, sometimes – to speed up – also at slightly elevated temperatures (up to 37-40 degr. C, 2-3 hrs) and sometimes also combined elevated temp. & under pressure (automatic devices). For evaluating tissue damage you should know first how to approach, and esp. choosing a sectioning plane through the organ you study which will show the matter you are after in a correct manner (anatomical issue). In the case of human heart or human kidney or human lung it is worth to know also that section dimension could be as large as 5-10 cm in diameter (and more): so you have to choose again which area you would like to evaluate or at least how you’ll divide the gross portions of your organs correctly (but if it is mice – what we don’t know for now – or rat or, or…? this might be no problem at all). FIXATION TIME (what is too short, what is o. k, what is too long?) in the range of at least 6 hrs RT (room temperature, use of a specimen probe rotator device) for a block say 10:10:5 mm I.M.H.O. as a minimum perhaps is ok ; immerging the spec. block too short produces a block unfixed in the center portion and therefore you will not be able to get a section of good quality. I agree with Wayne Briner in using always fixative 20-25 x the volume of the specimen. Tissue rotated during fixation in a specimen rotator device will be fixed easier and more thoroughly than tissue which just lies at the bottom of a phiole / sample glass tube . STORAGE in fixation solution MIGHT be no problem for ONLY morphological evaluation of tissue damage, but there are some CAVEATS: formaldehyde solution has been reported to act as dehydrating agent, if you store them too long in the fixative your specimens could get too hard and therefore – when embedded afterwards e.g. in Paraffin – the fixed tissue gets brittle properties which makes sectioning difficult, especially also if you use too long dehydration steps (We have not heard/read about dehydration or embedding issues (alcohols, or acetone, Paraffin or cryo-techniques so it would be interesting to get knowledge about your thoughts). Formaldehyde in slightly alkaline conditions (= pH 7,2 to 7.4; and not only those) is known to polymerize slowly but consistently all the while… does harden the specimen block…. so it might be advisable to lengthen the washing steps (with an appropriate buffer solution) after fixation. Also, formaldehyde solution left with the tissue produces formic acid (one of some byproducts) which is neutralized by the used (thats why we have to buffer the FA-solution, otherwise you run into acid fixation which distorts your tissue). It is right to say that also ARCHIVAL SPECIMENS stored for years in buffered Formaldehyde solution have been processed into paraffin blocks successfully, but this might not be the rule or be useful either. In general storage up to one week (or 2, especially at 4 degrees C) is acceptable and has been reported often as to be not damaging tissue per se. SPECIFIC FIXATIVES for different organs: YES, but usually this is an issue if you do vascular perfusion of whole organs. Then it might be advisable to adjust the fixative (also with additional cationic substances like CaCl2, MgCl2, etc) for perfusing brain, another for kidney, and another one for liver…. Note: Storage of tissue after fixation in 0.01 M PBS, as suggested by Lorena Salvatore I would not recommend due to hypo-osmolal(r)ity (unless the used PBS here is 10x concentrated): the usual concentration of storage buffer I.M.H.O.(in my honest opinion) is 0.1 to 0.15 M (0.13 M PO4 = equals to 300 mOsmol = the osmolarity of blood). Concerning "...1 day before mue.." in your initial question: I don't know what you mean with this but it could be "before dehydration, before …??). I know that my lengthy note perhaps is not very welcome (I don’t want to proselyte someone here) If you have more specific answers or also specific questions, please don’t hesitate to send an e-mail to [email protected] [EDITED 2016-08-02: DELETE: [email protected]   since meanwhile retired:  SEND YOUR REQUEST if needed to [email protected], thank you!] or to post this in ResearchGate-Topic . Best wishes and good luck with your work, Wolfgang MUSS, PhD, EM-Lab of Univ. Inst. Pathology, SALZBURG, Austria-EUROPE

NB: [only e-mail address edited, see in text above:

EDITED 2016-08-02: DELETE: [email protected] since meanwhile retired: SEND YOUR REQUEST if needed to [email protected], thank you!] 

Good fixation is based on the tissue size, time, organ specialty, and method. Of course, the hear cycling perfusion is the best way to fix the tissues for animal specimen. For human tissues, the way is used by me as following: 1) cut the tissues that can show the biggest area , but less than 0.3 cm thickness, drop to the Buffered 10% Formalin (1:20, Tissue/Formalin value) immediately at RT for 6-8 hrs, then for overnight at 4C; 2) For lung tissue, it is always float on the surface of the solution, please use the container to hold them sinking in solution, 3) Fixative-time is based on the tissue types, 24 hours are good enough for tissues of your list.

Hi you can use 10% formaldehyde for fixing the invertebrate tissues, not more than 2 days or else u can use bouin's fixative for gud results for vertebrate tissues like rat kidney , heart, muscles , spleen , liver tissues for histology and stain using haemotixylin and eosin for good results

NBF (Neutral Buffered Formalin) is widely used for general tissue fixation.

Some hospitals use FS (Formol Saline), as it's cheaper, but adequate.

Provided the tissue is not thicker than 10mm, 24-48 hours in NBF is sufficient, then change to 70% alcohol before wax processing.

Over fixation may interfere with antigenic studies.

You need about 20X NBF to the the volume of tissue.

Although Neutral Buffered Formalin has been widely used for general tissue fixation but what I personally experienced that fixation of these tissues in alcoholic Bouin's fixative always gives best histological details but one should follow best practices during processing of tissues up to block formation. Main culprit in Neutral Buffered Formalin fixed tissues r clearing agent and temp. which generally hardened the tissues.

Well friends, the choice of fixative and time of fixation depends mainly upon the purpose like routine histology by HE, special staining or immunostaining or electron microscopy and so on. Yes for routine staining best fixative is 10% NBF or 10% aquaeous formalin in neutral D W with a fixation time lapse of 24-48 hours. But for glycogen or fat or urate staining special fixatives are required. The time of fixation will also depend upon the size the density and thicness of the tissue to be fixed. For immunostaining, Much shorter fixation of not more than 12 hours is required as a longer fixation may mask most , particularly surface antigens. Therefore, for routine lesion scoring, as asked orignally, formalin fixation is simple, universal and most widely acceptable technique, of course foregoing its deleterious effects for the user.

After NBF fixation,I always wash my tissues 5-6 hours under running tap water in tubes tightly closed with holes all around the tube. This is to remove formalin deposits. Than I put samples in 70% ethanol O/N at room temperature. The following morning I start my dehydration process and finally paraffin inclusion.

Tissue morphology is best preserved in 10% Neutral Buffered Formalin, even if you considered further and advanced testings such as immunohistochemistry (IHC) technique.

However, these have been repeatedly written above. But, one more important thing in my points here is that I would advise you don't prolong your specimens longer in the fixative; usually it is stated between 6-24 hours. This is important because some antigen epitopes cannot withstand the effects of formalin for long time despite retrieval process, there would still be effect on the antigen. Again, your antibody of choice was not stated here for cross examination, this is another topic entirely considering how individual antibody behaves and survives post-fixatively.